AN ABSTRACT OF THE THESIS OF
Angel Luis Saavedra for the degree of Master of Science in Botany and Plant Pathology
presented on March 29, 2006.
Title: Susceptibility of Golden Chinguapin (Chrvsolepis chrvsophylla) to Phvtophthora
cambivora.
Abstract approved:
Redacted for privacy
Everett M Hansen
In early 2000, unusual mortality of a native North American tree, golden chinquapin,
was reported by the USDA-Forest Service. Dying trees exhibited girdling cankers in the
inner bark of the lower bole, branch flagging and defoliation. Isolations from necrotic
tissues and soil associated with diseased or killed trees yielded Phytophthora cambivora,
a pathogen that is known to infect and kill chestnut species in Europe and in the United
States.
Morphological, physiological and molecular testing confirmed the identity of isolates
recovered as P. cambivora. Pairing tests showed that both mating types, Al and A2, of
this species were present in forest soil in Oregon.
Pathogenicity tests were conducted to confirm the susceptibility of golden chinquapin
to P. cambivora. Two inoculation trials were conducted: I) Trees were wound inoculated
with mycelial plugs of P. cambivora; after 35 days necrotic tissues were formed in the
inner bark of all the inoculated trees, as seen in natural infections. 2) Seedlings were
inoculated with a zoospore suspension of P. cambivora; after 38 days all inoculated
seedlings were killed, the roots were rotted and the inner bark of lower stems was
necrotic. Phytophthora cambivora was re-isolated from necrotic tissues in both trials,
completing Koch's postulates.
©Copyright by Angel Luis Saavedra
March 29, 2006
All Rights Reserved
Susceptibility of Golden Chinquapin (Chrysolepis chrysophylla) to Phytophthora
cambivora
by
Angel Luis Saavedra
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Master of Science
Presented March 29, 2006
Commencement June 2006
Master of Science thesis of Angel Luis Saavedra
presented on March 29, 2006.
APPROVED:
Redacted for privacy
Major Professor, represeniihg Botany and Plant Pathology
Redacted for privacy
Chair of the Depar,ent ootany and Plant Pathology
Redacted for privacy
Dean of
raIuate School
I understand that my thesis will become part of the permanent collection of Oregon State
University libraries. My signature below authorizes release of my thesis to any reader
upon request.
Redacted for privacy
Angel Luis Saavedra, Author
ACKNOWLEDGMENTS
I am deeply grateful to all the people from Everett's Lab... Wendy my sincere and
deep gratitude for all the time, knowledge and words of encouragement that you gave me
every time I needed them (or not). Paul you were also always there for me and ready to
give so many great ideas to go through and smoothly finish this project.
I would also like to thank Drs. Parke, Stone, Spatafora and Miller for all the time and
patience that you dedicated to me during my studies at Oregon State University.
To all the people from the Botany and Plant Pathology Department that were always
there for me.
The staff of the Dorena Forest Tree Improvement Center in Cottage Grove and the
Forest Health Plant Protection group in Central Point, especially Don and Ellen Goheen
(and their cats), thanks a bunch for all your guidance, and friendship that you always
provided to me.
Eunsung my great friend, thanks to your inspiration and advice that I always found on
you, this project is completed. Also to the innumerable graduate colleges that were there
to pick me up during my ups and downs; Susan, Dave, Nate, Heather, John B., thank you.
My mother Corina, my father and all my brothers Ivan, Beto and Alexis and to all the
rest of my family I wish to thank you (muchas gracias) for all your support and love.
Finally, I can express with words how deeply indebted I am to Dr. Everett Hansen;
your guidance, friendship and support made this project possible. Thank you Everett
from my heart I will always be grateful to you and you also can count on me.
TABLE OF CONTENTS
Chapter 1. Susceptibility of golden chinquapin to Phytophthora cambivora ......... 1
1.1.Theproblem ..................................................................... I
1.2. Phytophthora cambivora (Petri) Buisman ................................... 3
I .2a Description of P. cambivora ......................................... 3
1 .2b Infection and spread biology ............................................ 5
1 .2c Distribution of Phytophthora cambivora ........................... 6
1 .2d P. cambivora host range ............................................. 8
1 .2e Important diseases caused by P. cambivora ....................... 9
1.3 Chrysolepischrysophylla ....................................................... 11
l.3a Distribution-range ..................................................... 12
1 .3b Taxonomy .............................................................. 12
1 .3c Diseases affecting golden chinquapin ................................ 13
1.3d Ecological roles and uses ............................................. 14
1.4 Thesis objectives ................................................................. 15
Literaturecited ................................................................................. 15
Chapter 2. Identification and pathogenicity of a Phytophthora species isolated from
dying golden chinquapin trees and Oregon forest soil ........................... 20
2.1 Introduction ....................................................................... 20
2.2 Material and methods ............................................................ 23
2.2a Isolation methods ...................................................... 23
TABLE OF CONTENTS (Continued)
Page
2.2b Isolate identification ................................................... 24
2.2c Pathogenicity test ...................................................... 26
Tree inoculation trial ............................................... 26
Seedling inoculation trial .......................................... 27
2.3 Results ............................................................................. 29
2.3a Isolate identification ................................................... 29
Colony description ................................................. 29
Mating type testing ................................................ 30
Sporangia description .............................................. 33
Optimal growth temperature...................................... 33
ITS rDNA sequencing ............................................. 35
2.3b Pathogenicity testing .................................................. 35
Tree inoculation trial ............................................... 35
Seedling inoculation trial .......................................... 40
2.4 Discussion ........................................................................ 42
Literaturecited ................................................................................. 47
Chapter 3. Summary and Conclusion ...................................................... 51
Bibliography ................................................................................... 55
Appendices ..................................................................................... 63
Appendix A. rDNASequence alignments of chinquapin isolates and
TABLE OF CONTENTS (Continued)
isolate AG45 clone 2 ............................................... 65
Appendix B. Pictures of necrotic lesions on ten golden chinquapin
trees inoculated with P. cambivora isolates ..................... 66
Appendix C. Pictures of golden chinquapin seedlings before and
after inoculation with P. cambivora ....................................... 77
LIST OF FIGURES
Page
2.1. Colony morphology of isolates of Phytophthora cambivora at room
temperature on carrot media after 4 days .......................................... 30
2.2. Amphigynous antheridia and oogonia of isolates of
Phytophthora cambivora ............................................................ 32
2.3. Mean daily growth rate (mm) from three plates per 3 isolates of P. cambivora
tested in this study .................................................................... 34
2.4. Bark lesions on bole of golden chinquapin trees .................................... 36
2.5. Basal lesion area (cm2) by tree per isolate of P. cambivora inoculated
in the bark often golden chinquapin trees ........................................ 38
2.6. Mean lesion area on ten boles of golden chinquapin inoculated with three
isolates of Phytophthora cambivora ............................................... 39
2.7. Mean lesion area (cm2) of 3 isolates per ten trees ................................... 39
2.8. One-year-old golden chinquapin seedlings 38 days after inoculation
with isolates of P. cambivora under greenhouse conditions ................... 41
2.9. Root conditions of one-year-old golden chinquapin seedlings 38 days
after inoculation with isolates of P. cambivora growing under
greenhouse conditions ............................................................... 41
LIST OF TABLES
Page
1. Geographical distribution of Phytophthora cambivora ............................... 6
2.1. Isolates of Phytophthora spp. collected and used in this research................. 24
2.2. List of trees and shrubs of golden chinquapin and locations were fruits
were collected ........................................................................ 27
2.3. Comparison of the characters of the isolates used in this study and
those reported in the literature ...................................................... 31
2.4. Comparison of growth rate among isolates of P. cambivora in this study....... 34
2.5. Basal lesion area (cm2) by tree per isolate of P. cambivora inoculated
in the bark often golden chinquapin trees........................................ 37
2.6. Analysis of variance for isolates effect (including the control as an isolate)
on mean lesion area in inoculated boles of Chrysolepis chrysophylla ........ 38
2.7. Analysis of variance for isolates effect on mean lesion area on inoculated
boles of Chrysolepis chrysophylla with isolates of P. cambivora
(control not included) ................................................................ 39
2.8. Analysis of variance for tree effect on mean lesion area in boles of
Chrysolepis chrysophylla inoculated with isolates of P. cambivora
(control not included) ................................................................ 40
2.9. Mortality of golden chinquapin inoculated with isolates of P. cambivora ....... 41
Susceptibility of Golden Chinquapin to Pliytophthora cambivora (Petri) Buisman
Chapter 1
Introduction
1.1 The problem
A new canker disease was reported in recent years (Goheen pers. comm.) causing
mortality of golden chinquapin trees (Chrysolepis chrysophylla (Dougi.) Hjelmqvist)
growing in the southwestern Oregon Cascade Range in the Butte Falls Ranger District of
the Rogue River-Siskiyou National Forest. The cankers on the boles of golden
chinquapin trees look similar to those that are caused by species of Phytophthora
(Oomycetes) on other species of North American forest trees (Tainter et al. 2000; Rizzo
et al. 2002; Betlejewski et al. 2003). The cankers in the inner bark were reddish-orange
in color extending upward from infected roots. Leaf flagging was a commonly observed
symptom of infected trees, especially on the lower branches. The leaves of affected trees
turned from their normal green to a bright red, almost like the tree is on fire, and
eventually were dropped. Preliminary surveys showed that most of the symptomatic or
dead trees were located near roads, although in some cases individual trees located
further away from roads were symptomatic.
This area of southwestern Oregon where symptomatic chinquapins have been observed
is approximately 28 miles east of Medford, OR on State Highway 140 and at
2
approximately 1 300m above sea level. The site's geographic co-ordinates are 42° 23'
north latitude and 122° 22' west longitude. In this area, golden chinquapin is associated
with a dominant coniferous forest that includes Douglas-fir (Pseudotsuga menziesii),
western white pine (Pinus monticola), western hemlock (Tsuga heterophylla), white fir
(Abies concolor), and Pacific yew (Taxus brevifolia). Other common plant species
associated with this forest community include Pacific madrone (Arbutus menziesii),
California (western) hazel (Corylus corn uta var. calfornica), Pacific dogwood (Cornus
nuttallii), Pacific rhododendron (Rhododendron macrophyllum), and Oregon grape
(Berberis nervosa).
In order to determine a possible causal agent of this new canker disease, samples of
bark from the actively growing canker margin were collected and plated in Phytophthora
selective medium (Hansen and Goheen, pers. comm.). Isolates of a Phytophthora species
were readily obtained from the cankers, suggesting that this organism could be the causal
agent of the disease. To further investigate the distribution of this organism in the area,
soils samples were collected from the bases of dying trees and baited using standard
techniques as described by Erwin and Ribeiro (1996). A Phytophthora species was
consistently recovered supporting suspicions that a soilborne Phytophthora was infecting
golden chinquapin through the roots and causing the lethal cankers.
Because isolates from soil and from cankers on dying golden chinquapin were similar
in morphology to P. cambivora, and also because this species is well known in Europe as
the causal agent of root rot and collar rot on commercially and ecologically important
hardwood species such as Juglans regia, Fagus sylvatica and Castanea sativa (Delatour,
2001; Jung et al. 2003; Vettraino et al. 2003), it was hypothesized that P. cambivora
was the causal agent of this new canker disease of golden chinquapin.
The objectives of this study were to identify the isolates recovered from dying
chinquapin and surrounding soil by morphological and molecular comparison with
known Phytophthora species, and to determine their association with this new canker
disease by satisfying Koch's postulates. Pathogenicity tests on golden chinquapin
seedlings under greenhouse conditions as well as inoculation in bark of healthy
chinquapin trees in the field were completed.
1.2 Phytophthora cambivora (Petri) Buisman
1 .2a. Description of P. cambivora
Petri (1917) in his research on root rot of chestnut in Europe described this species for
the first time under the name of Blepharospora cambivora Petri. Ten years later,
Christine Buisman (1927), as part of her thesis, argued that the characters on which the
genus Blepharospora was founded could not be considered distinct from the genus
Phytophthora and so in 1927 this species was synonymized under Phytophthora.
Members of the genus Phytophthora absorb nutrients from the environment, have
filamentous mycelia, and reproduce by spores. They share these features with true fungi,
however, the genus Phytophthora possesses zoospores with two flagellae and the
absorptive hyphae are diploid, among several other important differences. Current
taxonomic classifications place this genus under the Kingdom Chromista, within the
family Pythiaceae of the order Peronosporales (Cavalier-Smith, 1986; Kendrick, 2000).
4
Under this classification, the genus Phytophthora is more closely related to brown algae
than true fungi (Eumycota).
The sporangia ofF. cambivora are non-caduceus and non-papillate and their
antheridia are amphigynous. On the basis of these morphological features, Waterhouse
(1963) classified Phytophthora cambivora under group VI which includes another
important plant pathogen, Phytophthora cinnamomi. Morphologically, these two species
differ because Phytophthora cambivora produces bullate oogonia and no
chiamydospores. It should be noted that the Waterhouse morphological grouping of the
genus Phytophthora does not necessarily represent phylogenetic relationships, as was
demonstrated by Cooke et al. (2000). Under the latter phylogenetic scheme,
Phytophthora cambivora was placed in the same dade as Phytophthora cinnamomi.
Interspecific hybridization is a concept that has been proposed to explain pathogenicity
pattern changes on the genus Phytophthora (Brasier, 2000; May et al. 2003). It
hypothesizes that progeny resulting from interspecific hybridization could exhibit
reduced aggressiveness toward the parental hosts or host ranges that differ from those of
parental species. The latter is the case with a recently described new species of
Phytophthora (P. alni) in Europe affecting trees of the genus Alnus (Brasier et al. 2004).
The parent species are apparently P. cambivora and a species of Phytophthora related to
P. fragariae. It is also important to note that P. cambivora is a heterothallic species,
meaning that it requires a compatible opposite mating type to complete sexual
reproduction.
1.2b. Infection and spread biology
Most members of the genus Phytophthora are plant pathogens and P. cambivora is not
an exception. It is known that P. cambivora is a soil born pathogen and the lack of
chlamydospores suggests that the primary propagules of this pathogen consist of
swimming zoospores. In areas where compatible mating types are found and oospores
are formed, it can be inferred that these structures also can constitute a source of
inoculum. It can also be inferred that propagules of P. cambivora can be carried around
in the soil by many vectors including human, and that it may be distributed in from
infested areas to non-infested ones. Infection is presumed to occur through the roots of
trees, since it has been observed that the biomass of roots growing in soil free of
Phytophthora inoculum is greater than the mass of roots growing in infested soil (e.g.
Quercus spp) (Delatour, 2001) and the pathogen causes root rot on Prunus spp (Mircetich
and Matheron, 1976) and on English walnut (Vettraino et al. 2003). The results of the
latter study showed that P. cambivora, as well as other species of Phyt op ht ho ra
significantly reduced the biomass of roots of walnut seedlings growing in infested soil
with Phytophthora compared to uninoculated control roots. This suggests that this
pathogen's primary way of infecting its host is through the roots.
Whether P. cambivora can penetrate host tissues by other means than through their
roots is not known. Artificial inoculations of recently cut logs of conifer and hardwood
species (Hansen and Sutton, 2002) as well as seedling stem inoculations show that, given
the proper conditions, P. cambivora can successfully infect trees through wounds
(Delatour, 2001 and personal experience that will be discussed in Chapter 2).
Another mechanism of spread is by means of infected rootstocks as was documented
by Jeffers and Aldwinckle (1988). In this study, unbudded apple rootstocks from
shipments received by local nurseries from the major rootstock suppliers were sampled
before they were planted. Bioassay techniques included washing the rootstocks and
plating the rhizosphere soil on PAR (pimaricin, ampicilin and rifampicin) and PARH
(PAR plus Hymexazol) selective media, as well as baiting the soil using apple cotyledons
as baits. They recovered Phytophthora cambivora 111 times from 153 roots of unbudded
clonal apple rootstocks. They also reported that most recovered isolates of P. cambivora
were mating type Al (92 isolates) and 3 isolates of mating type A2. This suggests that
opportunities for intraspecific sexual reproduction, and hence oospores as a source of
host infection, were reduced.
1.2c. Distribution of Phytophthora cambivora
According to the Commonwealth Mycological Institute (CMI) Description of
Pathogenic Fungi and Bacteria No. 112 (Waterhouse and Waterston, 1966),
Phytophthora cambivora has a worldwide distribution (Table 1).
Table 1. Geographical distribution of Phytophthora cambivora
Continent
Africa
Asia
Australia
Europe
& Oceania
North America
Countries
Mauritius
India
New Zealand
Azores, France, Great Britain, Italy,
Poland, Portugal, Spain, Switzerland,
Turkey and Yugoslavia
Canada and the United States
7
Since this publication, P. cambivora has been reported from many other countries
including South Africa, Japan, Australia and Scotland and is associated with diseases of
endemic trees as well as ornamental crops (Gerretson-Cornell 1978; Erwin and Ribeiro,
1996). Several researchers during the past few years have studied the involvement of
Phytophthora species with a complex disease known as "oak decline" in Europe.
Vettraino et al. (2002) reported recovering P. cambivora and eleven other Phytophthora
species, from sites associated with oak decline syndrome in the southern and central part
of Italy and with chestnut in the Italian northern region. Jung et al. (2000) investigating
the distribution of Phytophthora species from different oak stands in Bavaria, Germany
recovered P. cambivora from soil. Researchers from other countries such as England and
Sweden investigating the matter have recovered P. cambivora from oak stands (Brasier
and Jung, 2001; Jönsson et al. 2003 respectively).
Phytophthora cambivora is also associated with another important disease known as
"ink disease" in Europe and in the USA. Studying the distribution of the disease in
chestnut stands in Europe, Vettraino et al. (2005) reported recovering isolates of P.
cambivora from chestnut in Greece, Italy and France.
The first report of the occurrence of P. cambivora in the United States was by Pirone
(1940). He reported mortality of hundreds of maple trees in New Jersey due to a basal
canker associated with the P. cambivora group. Other studies have reported the
occurrence of P. cambivora in California and New York, always associated with diseases
affecting fruit orchards (Mircetich et al. 1974; Mircetich and Matheron, 1976; Jeffers and
Aldwinck!e, 1988). In the Pacific Northwest, P. cambivora has been isolated from dead
E;1
roots and girdling stem cankers on Noble fir in Christmas tree plantations (Hamm and
Hansen, 1987; Chastagner et al. 1995).
This pathogen has also been reported from New Guinea (Ash, 1988). In this study,
searching for explanations of the observed dieback of Nothofagus, P. cambivora along
with P. cinnamomi Rands were isolated from soil samples in Nothofagus forest in New
Guinea. Both Phytophthora species were isolated from healthy and declining stands and
no conclusion could be inferred regarding pathogenicity on Not hofa gus.
More recently, Greslebin et al. (2005) isolated P. cambivora from soil associated with
Austrocedrus chilensis forest in Patagonia, Argentina. It was recovered from soil in only
one of eleven sites of the study; whether this pathogen is associated with the observed
decline of A. chilensis is still not known.
I n summary, P. cambivora has a worldwide distribution and because it has been found
associated with healthy, declining, and dead stands it is likely that this pathogen occurs in
many other places yet to be described.
1.2d. P. cambivora host range
Erwin and Ribeiro (1996) reported a list of hosts susceptible to P. cambivora that
includes more than 30 species in 12 families of plants. Depending upon the host, this
pathogen causes diseases that include wilt, blight, collar rot, root rot and trunk canker.
Most of the hosts included in the list are of economic and ecological importance; among
others are species of the genera Acer, Castanea, Casuarina, Fagus, Juglans, Malus,
Nothofagus, Persea, Prunus, Rhododendron, Rubus and Ulmus. The list also includes
members of the Asteraceae family that are cultivated for medicinal purposes such as
Chrysanthemum cinerariafolium and nursery plants like species of the genus Senecio.
Phytophthora cambivora has been isolated from a number of species of conifers grown
for Christmas trees in the Pacific Northwest of the United States. Hamm and Hansen
(1987) reported isolating P. cambivora from rotted roots and stem cankers on 2-5 year-
old Noble fir but pathogenicity was not confirmed at the time. Chastagner et al. (1990a)
and Chastagner et al. (1995) also reported isolating this pathogen, among other species of
Phytophthora, from stem cankers and root rot of symptomatic Noble fir trees in
plantations in Oregon and Washington but did not test whether P. cambivora could cause
root rot and stem canker and consequently mortality on Noble fir. However, Chastagner
et al. (1990b) reported a pathogenicity test in which stems of field grown Noble fir were
inoculated with species of Phytophthora, including P. cambivora, in order to study their
ability to cause canker. The results indicated that P. cambivora was among the most
virulent species of Phytophthora causing stem canker in Noble fir. In this same report,
eleven species of Abies were grown in soil infested with several species of Phytophthora
in order to determine susceptibility of these hosts to these pathogens. Phytophthora
cambivora was found to be virulent to susceptible species of Abies but P. cinnamomi
caused the most mortality.
1.2e. Important diseases caused by P. cambivora
Among the diseases caused by P. cambivora, the ink disease of chestnut in Europe and
in the United States is considered to be the better known and studied disease (Erwin and
Ribeiro, 1996; Vettraino et al. 1999; Bourbos and Metzidakis, 2000). This disease is
10
characterized by trunk and root rot and consequently tree mortality. Another feature of
this disease is the occasional exudation of inky fluid from dying or dead bark at the base
of the trunk. Little is known about how this pathogen is moved around in nature.
However, it is likely that it is carried in mud by humans and animals as noted by
Vettraino et al. (1999). In addition, this report states that in sweet chestnut plantations in
Italy, the disease commonly starts from trees along roads and trails. This pattern has also
been observed on cankered golden chinquapin trees, as will be discussed in chapter 2 of
this dissertation.
Due to the economic importance of sweet chestnut (estimated to have an import
wholesale value approaching $20 million in Italy alone) and the fact that great concern is
apparent in Europe regarding a potential new outbreak, a considerable amount of funding
for research is being put into place in search of a better understanding of this disease. It
has been estimated that annual consumption of chestnut per capita is around 2 lb. in
China, 1 lb. per capita in Europe and less than half a pound per capita in the U.S.
Another reason that chestnut ink disease receives so much attention is to try and avoid
ecological catastrophes like the one caused by Cryphonectria parasitica on American
chestnut early in the 20th century in the United States.
Other diseases of economic importance caused by P. cambivora are crown and root rot
of many fruit trees such as plums, apples, apricot, peaches and cherry among others
(Mircetich and Matheron, 1976; Jeffers and Aldwinckle, 1988; Erwin and Ribeiro 1996).
Unfortunately, above ground symptoms of Phytophthora infected trees resemble those
caused by other root rot pathogens, so direct isolation from infected plant tissues is
necessary to determine whether an infection has been caused by P. cambivora.
11
Phytophthora cambivora, as well as other species of Phytophthora, has been
associated with oak decline in Central Europe (Jung et al. 2000). In this study,
Phytophthora cambivora was among the most frequently recovered species, but only
from soil samples. Whether there is a correlation between the occurrences of P.
cambivora in the soil and the severity of oak decline is still undetermined. It was noted
in this study that characteristics of the site (soil pH, drainage, texture) were strongly
related to the presence of Phytophthora species.
1.3. Chrysolepis chiysophylla
Golden chinquapin (Chrysolepis chrysophylla), also known as giant chinquapin, is a
hardwood species that grows in a landscape dominated by coniferous forests. In Oregon
and northern California, golden chinquapin grows in tree and shrub forms in a variety of
habitats but it is rarely a dominant component of any stand. The tree form can be found
growing as stem clumps or as solitary trees. Toward its southern range, trees commonly
grow as tall as 31 m and reach circumference of 120 cm in diameter (Jensen and Ross,
1995). Golden chinquapin is an evergreen species; its leaves are arranged in an alternate
pattern on the branches and they are coated underneath with golden yellow leaf-hair,
hence its common name. Even though its wood posseses excellent qualities for furniture
making and for hardwood plywood, this species is considered in many areas of its natural
range to be a competitor of commercial species.
12
1 .3a. Distribution-range
Mc Kee (1990) reported that the natural range of golden chinquapin extends from San
Luis Obispo County in California, to Mason County in western Washington. He noted
that in California, golden chinquapin grows primarily in the Coast Ranges, but there is a
disjunct population in El Dorado County in the Sierra Nevada. In Oregon, it is found in
the Coast Ranges as far north as Benton County, and throughout the Cascade Range as
far north as Marion County (pers. observations). In Washington, golden chinquapin is
represented by two disjunct populations in Mason and Skamania Counties (Kruckeberg,
1980).
Shrub forms of the species are found throughout its range. The tree form is primarily
distributed from Lane County, OR, south to Mann County, CA. It is found from near sea
level in the Coast Ranges of Oregon and California to over 1525 m in elevation in the
Cascades. Although golden chinquapin is generally thought of as a mid- to low-elevation
species, the shrub form can be found along the crest of the Cascade Range in Oregon
from 1525 to 1830 m.
1.3b. Taxonomy
The species
Chrysolepis chrysophylla
(Dougl. ex Hook.) Hjelmqvist is classified under
the botanical family of Fagaceae. Members of this family are characterized as catkinbearing trees and shrubs with the fruit, an acorn, at least partially enclosed by a cupule
(Smith, 1977).
Golden chinquapin staminate flowers are creamy white and arranged in catkin-like
inflorescences. The flower odors that are perceived during the summer as it blooms are
13
distinctive of this genus. The female flowers are borne within an involucre at the base
of male flowers or can be located alone along the stem. The nuts, which mature in the
fall of the second growing season, are enclosed in a spiny burr that is unique for this
genus (Manos et al. 2001).
The genus Chrysolepis includes only two species and both are native to the western part
of the United States; C. chrysophylla and C. sempervirens. The separation of this genus
from the previously known genus Castanopsis occurred in 1948 on the basis of the
cupule structure (Hjelmqvist, 1948). The genus Castanopsis was reserved for the
subtropical and tropical species that occurred in Asia.
Finally, naturally occurring and cultivated species of "chinquapin" in the SE United
States are members of the genus Castanea. According to Li et al. (2004), Castanea,
Castanopsis and Chrysolepis are distinct but closely related genera.
1.3c. Diseases affecting golden chinquapin
There are few reports of diseases or insects that affect the growth or survival of golden
chinquapin. (Hepting, 1971). However, it is known that it is very susceptible to a number
of root rot pathogens, heart rots, and a few common foliar fungi (Fan et al. 1989).
Among these organisms heart rot caused by Phellinus igniarius is known to be damaging
to golden chinquapin, as well as root rot caused by Arm illaria species. During the current
study infection by Arm illaria on golden chinquapin trees was observed at different sites
in its natural range. However, no attempt was made to identify the species of Armillaria.
14
There are very few insect pests on golden chinquapin. Seed-feeding insects are the
most commonly reported. These insects could play a significant role in reducing golden
chinquapin trees' regeneration capacity depending on the percentage of infested fruits.
Dryocosmus castanopsidis,
a cynipid wasp attacks the staminate flowers causing
spherical golden-yellow to brown galls (Goheen pers. comm.). The amount of infested
flowers also varied by region and from tree to tree. No study has looked into its effects
on golden chinquapin reproduction.
l.3d. Ecological role and uses
Although, golden chinquapin has a light brown and fine grained wood, it has little
commercial value because trees of timber size are not abundant at any one place
(Brockman, 1958; Jensen and Ross, 1995). It is known that wood of golden chinquapin
was used for making agricultural tools and several other items that required strong wood
(Uphof, 1959).
Ecologically, golden chinquapin trees are considered of importance for wildlife. Its nuts
are nutritious and eaten by a variety of birds and small mammals. The restriction is the
availability of fruits, since they are produced at irregular intervals and in low numbers
(Mc Kee, 1990).
Golden chinquapin shrubs are rarely browsed by livestock, but certain shrubby ecotypes
are of moderate importance as mule deer browse in portions of California. However,
golden chinquapin bushes are browsed only when the preferred browsing species is
unavailable due to overgrazing or other reasons (Hubbard, 1974). Historically, golden
15
chinquapin nuts were roasted and eaten by indigenous people throughout the Coast
Ranges of northern California and southwestern Oregon (Uphof, 1959).
1.4. Thesis objectives
I want to establish the identity of the Phytophthora isolates recovered from dying
chinquapin trees and surrounding soil and to determine their association with this new
bole canker on golden chinquapin by completing Koch's postulates. To accomplish these
objectives, the following activities were undertaken:
1).
Three Oregon isolates resembling P. cambivora were characterized in terms of their
mating type, ITS rDNA sequence, and morphological and physiological features.
2). Field inoculation of live trees with the recovered isolates resembling P. cambivora
were made to observe necrotic lesion formation, possible differences in aggressiveness
among them, and their re-isolation to complete Koch's rules.
3).
Seedlings of golden chinquapin were inoculated under greenhouse conditions with
the recovered isolates resembling P. cambivora to demonstrate susceptibility of seedlings
to zoospores of this pathogen. Mortality was measured, further satisfying Koch's
Postulates.
Our hypothesis is that the recovered isolates belong to P. cambivora and that this
species is the causal agent of this new canker disease reported in boles of golden
chinquapin in Oregon.
Literature Cited:
Ash, J. (1988). Nothofagus (Fagaceae) forest on Mt. Giluwe, New Guinea. New Zealand
Journal of Botany 26: 245 - 258.
16
Betlejewki, F., Casavan, K., Dawson, A., Goheen, D., Mastrofini, K., Rose, D. and
White, D. (2003). A Range-Wide Assesment of Port-Orford-cedar
(Chamaecyparis lawsoniana) on Federal Lands. Bureau of Land Management,
USDA- Forest Services. l82p.
Bourbos, V. and Metzidakis, I. (2000). Biological control of Phytophthora cambivora
(Petri) Buisman in chestnut tree (Castanea sativa Mill). Ecologia Mediterranea 26
(1-2): 123-127.
Brasier, C. M. (2000). The rise of the hybrid fungi. Nature 405: 134-135.
Brasier, C. M. and Jung, T. (2001). Progress in understanding Phytophthora diseases of
trees in Europe. In: Phytophthora in Forests and Natural Ecosystems. 2'
International IIJFRO Working Party 7.02.09 Meeting. Albany. W. Australia 30th
Sept. -5 Oct 2001. McComb, J., Hardy, G. and Tommerup, I. (eds.) (Murdock
University Print) Pp: 4-18.
Brasier, C. M., Kirk, S., Delcan, J., Cooke, D., Jung, T. and Man In't Veld, W. A. (2004).
Phytophthora alni sp. nov. and its variants: designation of emerging heteroploid
hybrid pathogens spreading on Alnus trees. Mycol. Res. 108(10): Pp: 1172 1184.
Brockman, C. F. (1958). Golden chinquapin. University of Washington Arboretum
Bulletin 21: 46-47, 70
Buisman, C. J. (1927). Root rot caused by Phycomycetes, University of Utrecht. Meded.
Phytopathol. Lab. Willie Commenlin Scholten 11:7. (Rev. Appi. Mycol. 1927,
6:380): 5lp.
Cavalier-Smith, T. (1986). The Kingdom Chromista: Origin and Systematics. j
Progress in Phycological Research. Round and Chapman (Eds.). Bristol, UK,
Biopress. Vol. 4: 309 - 347.
Chastagner, G., Hamm, P. and Byther, R. (1990a). Symptomology of Phytophthora root
rot and stem canker disease of Noble fir in the Pacific Northwest. Phytopathology
80(9): 887.
Chastagner, G., Riley, K. and Hamm, P. (1990b). Susceptibility of Abies spp. to seven
Phytophthora spp. Phytopathology 80(9): 887.
Chastagner, G., Hamm, P. and Riley, K. (1995). Symptoms and Phytophthora spp.
associated with root rot and stem canker of Noble fir Christmas trees in the
Pacific Northwest. Plant Disease 79(3): 290-293.
17
Cooke, D. B. L., Drenth, A., Duncan, J. M., Wagels, G. and Brasier, C. M. (2000). A
molecular phylogeny of Phytophthora and related oomycetes. Fung. Gen. Biol.
30: 17-32.
Delatour, C. (2001). Phytophthoras and oaks in Europe. In: Proceedings of the 49th
Annual Western International Forest Disease Work Conference. September 1014. 2001. Carmel Mission Inn, Carmel, CA. Pp:16 -30.
Erwin, D. and Ribeiro, 0. (1996). Phytophthora Diseases Worldwide. St. Paul,
Minnesota. USA., APS Press. 562p.
Fan, D., Bills, G., Chamuris, G. and Rossman, A. (1989). Fungi on Plants and Plant
Products in the United States., The American Phytopathological Society. St. Paul,
MN. USA. 1252 p.
Gerrettson-Cornell, L. (1978). Notes on the morphology of an isolate of Phytophthora
cambivora (Petri) Buis. from Australia. Phyton 36(1): 85 - 89.
Greslebin, A. G., Hansen, E. M., Winton, L. M. and Rajchenberg, M. (2005).
Phytophthora species from declining Austrocedrus chilensis forest in Patagonia,
Argentina. Mycologia 97(1): 218 - 228.
Hamm, P. B. and Hansen, E. (1987). Identification of Phytophthora spp. known to attack
conifers in the Pacific Northwest. Northwest Science 61(2): 103-109.
Hansen, E. M. and Sutton, W. (2002). Log inoculations to asses tree susceptibility to
sudden oak death. Phytopathology 92 (6 Supplement): S33.
Hepting, G. (1971). Diseases of Forest and Shade Trees of the United States. USDA
Agriculture Handbook No. 386. 658p.
Hjelmqvist, H. (1948). Studies on the floral morphology and phylogeny of the
Amentiferae. Bot. Not. Suppl. 2(1): l'71p.
Hubbard, R. L. (1974). Castanopsis (D.Don) Spach, chinkapin. In: Seeds of Woody
Plants in the United States. Schopmeyer tech. coord. Washington D.C., USDA,
Agriculture Handbook 450: 276-277.
Jeffers, S. N. and Aldwinckle, H. S. (1988). Phytophthora crown rot of apple trees:
Sources of Phytophthora cactarum and P. cambivora as primary inoculum.
Phytopathology 78: 328 - 325.
Jensen, B. and Ross, C. (1995). Trees to Know in Oregon, Extension Circular 1450.
Oregon State University Extension Services and Oregon Department of Forestry.
l28p.
18
Jönsson, U., Lunderberg, L., Sonesson, K. and Jung, T. (2003). First record of
soilborne Phytophthora species in Swedish oak forest. For. Path. 33: 175-179.
Jung, T., Blaschke, H. and O3wald, W. (2000). Involvement of soilborne Phytophthora
species in Central European oak decline and the effect of site factors on the
disease. Plant Pathology 49: 706-718.
Jung, T., Nechwatl, J., Cooke, D. E., Hartmann, G., Blaschke, H., OJ3wald, W. F.,
Duncan, J. M. and Delatour, C. (2003). Phytophthora pseudosyringae sp. nov., a
new species causing root and collar rot of deciduous tree species in Europe.
Mycol. Res. 107(7): 772 - 789.
Kendrick, B. (2000). The Fifth Kingdom. Newburyport, MA. USA, Focus Publishing. R.
Pullins Co. 173p.
Kruckeberg, A. R. (1980). Golden Chinquapin (Chrysolepis chrysophylla) in Washington
State: A species at the Northern Limit of its Range. Northwest Science 54(1): 416.
Li, R., Chen, Z., Lu, A., Soltis, D., Soltis P., and Manos, P. (2004). Phylogenetic
relationship in Fagales based on DNA sequences from three genomes. mt. J. Plant
Sci.165(2): 311-324.
Manos, P., Zhou, Z. and Cannon, C. (2001). Systematics of Fagaceae: phylogenetic test
of reproductive trait evolution. Tnt. J. Plant Sci. 162(6): 136 1-1379.
May, K. J., Drenth, A. and frwin, J. A. G. (2003). Interspecific hybrids between the
homothallic Phytophthora sojae and Phytophthora vignae. Australasian Plant
Pathology 32: 353-359.
Mc Kee, A. (1990). Castanoposis chiysophylla (Dougl-)A.DC. Giant Chinkapin. Jii
Silvics of North America, Burns, R. and Honkala, B. (eds). USDA-Agricultural
Handbook No.654. 877p.
Mircetich, S. M. and Matheron, M. E. (1976). Phytophthora root rot and crown rot of
cherry trees. Phytopathology 66: 549-558.
Mircetich, S. M., Matheron, M. E. and Schreader, W. R. (1974). Sweet cherry root rot
and trunk canker caused by Phytophthora spp. Proc. Am. Phytopathol. Soc.
(abstr.) 1: 58.
Petri, L. (1917). Ricerche sulla morfologia e biologia della Blepharospora cambivora,
parassita del Castagno. Atti. R. Accad. Lencei. Rend. Cl. Sci. Fis. Mat. Nat. Ser.
5 26: 297 -299.
Pirone, P. (1940). Phytophthora disease of maple. Phytopathology 30: 19.
Rizzo, D. M., Garbelotto, M., Davidson, J. M., Slaugther, G. W. and Koike, S. T. (2002).
Phytophthora ramorum as the cause of extensive mortality of Quercus spp. and
Lithocarpus densflorus in California. Plant Disease 86(3): 205 - 214.
Smith, J. P. (1977). Vascular Plant Families. Eureka, CA, Mad River Press, Inc. 320p.
Tantier, F., O'Brien, J. G., Hernandez, A., Orozco, F. and Rebolledo, 0. (2000).
Phytophthora cinnamomi as a cause of oak mortality in the state of Colima,
Mexico. Plant Disease 84: 394 -398.
Uphof, J.C.Th. (1959). Dictionary of Economic Plants. Weinheim, H.R. Engelmann
(J.Cramer); New York, Hafner. 400p.
Vettraino, A. M., Natili, G., Anselmi, N. and Vannini, A. (1999). Recent advances in
studies on Phytophthora species associated with Castanea sativa and Quercus
cerris in Italy. In: Phytophthora Diseases of Forest Trees. 1St International JUFRO
Working Party 7.02.09 Meeting. Corvallis, Oregon: Oregon State University.
USA. Hansen, E.M. and Sutton, W. (eds). Pp: 34-36.
Vettraino, A. M., Barzanti, G. P., Bianco, M. C., Ragazzi, A., Capretti, P., Paoletti, E.,
Luisi, N., Anselmi, N. and Vannini, A. (2002). Ocurrence of Phytophthora
species in oak stand in Italy and their association with declining oak trees."EQL
Path. 32: 19-28.
Vettraino, A. M., Belisario, M., Maccaroni, M. and Vannini, A. (2003). Evaluation of
root damage to English walnut caused by five Phytophthora species. Plant
Pathology 52: 491 495.
Vettraino, A. M., Morel, 0., Perlerou, C., Robin, C., Diamandis, S. and Vannini, A.
(2005). Ocurrence and distribution of Phytophthora species in European chestnuts
stands, and their association with ink disease and crown decline. Eur. J. For. Path.
111: 169-180.
Waterhouse, G. M. (1963). Key to the species of Phytophthora de Bary. Mycol. Pap. No.
92: 22p.
Waterhouse, G. M. and Waterston, J. M. (1966). C.M.I. Descriptions of Pathogenic Fungi
and Bacteria No. 112, Maps are released periodically by the Commonwealth
Mycological Institute (International Mycological Institute, Bakeham Lane,
Egham, Surrey, United Kingdom)
20
Identification and pathogenicity of a Phytophihora species isolated from dying
Golden Chinquapin trees and Oregon forest soil
Chapter 2
2.1 Introduction
Golden chinquapin (Chrysolepis chrysophylla (Doug!) Hjelqvist) is an evergreen tree
species native to western North America. It grows in a variety of woody plant
communities but it is most commonly found growing in mixed stands of conifers,
especially Douglas-fir, since golden chinquapin rarely occurs in pure stands. Its native
range extends from California (San Luis Obispo County) to Washington (Mason County).
It can be found growing in both the California and Oregon Coast Ranges and in the
Oregon Cascade Range; however in Washington only two isolated populations have been
found (Mason and Skamania Counties) in several surveys conducted in the State
(Brockman, 1958; Kruckeberg, 1980). Golden chinquapin can be found growing from
mid to low elevation ranges but it also can be seen above 1 500 m, growing mainly in its
shrub form.
This cone-triangular shaped tree that commonly grows up to 30 m tall and 120 cm in
diameter, stands out within the mixed forest where it grows due to its distinctive oval to
lanceolate, alternate leaves. When trembled by the wind their beautiful underneath
golden scale is revealed, which gives the common name of "golden chinquapin". In
addition, a strong musky odor that its flowers emit during late spring and early summer
reveals the presence of golden chinquapin in mixed stands.
21
Golden chinquapin fruits are triangular nuts enclosed inside sharp spiny burrs
resembling those of chestnuts (Castanea spp). The nuts are edible, and used by wildlife.
Botanist David Douglas (1799-1834) (Cited by Scheffer, 1961) wrote that after he shot a
band-tailed pigeon and examined the bird's stomach contents he found chinquapin nuts.
The market for golden chinquapin wood is very small, even though the wood is
considered to have excellent qualities for furniture and for construction lumber (McKee,
1990). Use is limited because the wood is very difficult to cure, very few mills are
equipped to process it, and pure stands rarely occur in nature. Other species of
"chinquapin" (Castanea spp.) in the southeastern United States are cultivated for their
sweet edible nuts, and seedlings of these species, especially Castaneapumila are
available in nurseries (Payne et al. 1993). Unfortunately, this is not the case for golden
chinquapin, even though the nuts are edible and a small market exists for the wood.
Golden chinquapin has a poor rate of seed germination, ranging from 14 to 53 percent
(Hubbard, 1974).
There are not many reports of diseases or insects that might affect the growth or
survival of golden chinquapin (Fan et al. 1989). However, it is susceptible to root rot
caused by Arm illaria species, flower galls caused by Dryocosmus captanoisidis (Russo,
1979 and personal observations) and defoliation caused by an unidentified ascomycete
fungus reported by Kruckeberg (1980) in Washington. Unlike its close relatives,
Castanea spp. in Europe and in North America, golden chinquapin is not susceptible to
Cryphonectria parasitica (Anagnostakis, 1987; Barnard, 2000).
Observations of increasing mortality of golden chinquapin trees in southwestern
Oregon (Don Goheen, personal communication) were brought to the attention of the
22
scientific community in 2001 (Hansen, 2001). The problem was described as a new
lethal canker disease of golden chinquapin characterized by girdling basal cankers
extending upward from infected main roots. The leaves of affected trees turned bright red
and later dropped, with subsequent death of the tree. A Phytophthora species, tentatively
identified as P. cambivora, was isolated from bark lesions as well as from soil
surrounding dying trees. Phytophthora cambivora is a common pathogen on hardwood
trees in Europe (Santini et al. 2001; Brasier et al. 2004) and was reported killing maple
trees in eastern United States (Pirone, 1940; Crandall et al. 1945), but almost nothing is
known about P. cambivora on golden chinquapin or in western forests in general.
This study is the first to examine in detail the host-pathogen system of golden
chinquapin and this Phytophthora species. The primary objective of this study was to
confirm the identity of the suspected pathogen, and to experimentally demonstrate its
pathogenicity to golden chinquapin. Confirmation of the hypothesis that golden
chinquapin is susceptible to P. cambivora provides the first step to understanding the
epidemiology of this new canker disease for forest managers.
In order to test the hypothesis that golden chinquapin is susceptible to P. cambivora
experiments were performed to:
1) Confirm the identification of isolates recovered from forest soil around healthy and
diseased golden chinquapin trees and from infected bole tissue using morphological
comparisons as well as molecular approaches.
2) Complete Koch's postulates by inoculating live trees in the field as well as inoculating
seedlings under greenhouse conditions.
23
2.2 Materials and Methods
2.2a. Isolation methods
The three isolates used in this research were collected from 200 1-2002 in forested areas
within the range of golden chinquapin in Oregon. Two isolates were recovered from
forest soil collected at the base of recently killed or healthy chinquapin trees.
Approximately 100 g of soil were deposited into a metal baking pan and flooded with 1 L
distilled water. Each soil was then baited with 5, one-inch-long pieces of Port-Orford-
cedar (Chamaecyparis lawsoniana) foliage, and with one partially submerged pear. Soils
were baited for 5-6 days at room temperature (-25°C) in the dark. Necrotic foliage baits
and necrotic spots on pears were plated onto the Phytophthora selective medium CARP
(17 g I L cornmeal agar, with 200p.g m11 ampicilin, 10 jtg mY1 rifampicin and 10 j.ig mY'
pimaricin amended with 50 .tg mY' hymexezol). Growing colonies with morphology
resembling Phytophthora species were subcultured onto carrot agar (CA) (Kaosiri et al.
1978; Vettraino et al. 2000; Brasier and Kirk, 2004) and maintained at room temperature
for further identification and pathogenicity testing.
A third isolate was recovered from an active advancing lesion in the inner bark of a
fading golden chinquapin tree. Small pieces of necrotic tissue were plated on CARP+
and observed for developing colonies. Colonies looking like Phytophthora were then
transferred to carrot agar for further identification. Table 2.1 summarizes the origin of the
isolates.
24
Table 2.1. Isolates of Phytophthora spp. collected and used in this research
ISOLATE SOURCE OF
4048
GEOGRAPHIC
YEAR
ISOLATE
LOCATION
COLLECTED
Bark
Butte Falls, OR. USA
July, 2001
COLLECTED BY
E.M. Hansen, D. Goheen, A.
Saavedra
4074
Soil
Samtiam Hy., OR. USA
August, 2001
E.M. Hansen
09 17-2
Soil
Butte Falls, OR. USA
September, 2002
A. Saavedra
2.2b. Isolate identfl cation
Isolates were identified by comparing growing colonies and their reproductive
structures against known Phytophthora isolates and descriptions of Phytophthora species
reported in the literature (Newhook et al. 1978; Stamps Ct al. 1990; Erwin and Ribeiro,
1996). To promote sporangial formation, mycelial plugs, approximately 5 mm diameter,
were taken from actively growing colonies of each isolate growing on CA. Plugs were
placed into sterile 9 cm petri dishes and flooded with ten ml clarified V8 broth (200 ml of
clarified V8 juice diluted into 800 ml of deionized water, Zentmyer et al. 1976) and
incubated for two days at 18°C (Ito and Kudo, 1994). Each plate was rinsed 3-5 times
using deionized water and then flooded with 10 ml of soil extract water (1 g soil / 100 ml
deionized water) and incubated for approximately 60 hours.
To determine mating type, the three chinquapin isolates recovered in this research were
paired with known tester isolates of Phytophthora cambivora, (P31-A2 and P32-Al,
courtesy of Sabine Werres, Braunschweig, Germany, and PC-98-1116-A1 from Portland,
OR, courtesy of the OSU Plant Disease Clinic), and Phytophthora cinnamomi, (PC-98-
224- A2 from Gaston, OR, courtesy of the OSU Plant Disease Clinic, and P33-Al from
25
Sabine Werres in Germany). All isolates were grown in CA for five days in the dark at
room temperature before pairing. Mycelial plugs from the active growing margin of the
various isolates were paired in all combinations on CA in 9 cm petri dishes. Isolates were
placed 2 cm apart and incubated at room temperature in the dark for 7-8 days, with two
replicates of each pairing (Kellam and Zentmyer, 1986).
For further confirmation of the identities of the three chinquapin isolates, DNA from
the internal transcribed spacer (ITS) region of the nuclear ribosomal gene was extracted,
amplified, and sequenced. A 5 nmi dia. plug was taken from colonies growing on CA
and DNA was extracted as described in Winton and Hansen (2001).
PCR was performed in 5Op.l reactions (lx buffer, 200nM dNTP, 0.4 .tM ITS4 and ITS5
primers (White et al. 1990), 0.05 UI p.1 RedTaq DNA polymerase (Sigma St. Louis, MO)
and 2 p.1 template DNA). The reaction conditions were: 60 s at 94°C, 35 cycles of 60 s at
94°C, 60 s at 5 5°C, 60 s at 72°C and a final incubation for 7 mm at 72°C. The PCR
products were prepared for sequencing by adding O.5.tl EXOSAP-IT (USB Cleveland,
OH) and incubated overnight at room temperature (-22°C) followed by 15 mm at 80°C.
Direct sequencing of PCR products (ABI Prism BigDye Terminator Cycle Sequencing
Ready Reaction Kit, Applied Biosystems, Foster City, CA) was performed with primers
ITS4 and ITS5 and run on an ABI Prism 3100 Genetic Analyzer (Applied Biosystems).
Contigs were assembled and edited with the Staden (1996) software package. Edited
sequences were compared to sequences of the known species available at GenBank with
the BLAST search utility (Altschul et al. 1997)
For determining optimal growth temperatures, isolates were grown on CA in 9 cm petri
dishes for two days at room temperature in the dark. After this incubation period, the
plates were moved to incubators at temperatures ranging from 5°C to 35°C for 4 days.
Growth rate was determined by measuring colony radius along two lines intersecting at
right angles at the centre of the inoculum (Jung et al. 1999; Werres Ct al. 2001; Jung et al.
2002; De Cock and Lévesque, 2004). Colony margins were marked every two days and
the average daily radial growth calculated. Three replicate plates per isolate/temperature
were used for this test.
2.2c. Pathogenicity tests
Tree inoculation trial
A field inoculation trial was conducted on the Rogue River National Forest, Butte Falls
Ranger District, in an area where P. cambivora has been recovered from soil and from
dying golden chinquapin trees (Table 2.1). This area, located on Oregon State Highway
140 in southwestern Oregon between milepost 28 and milepost 30 has a larger number of
dead or dying golden chinquapin trees than any other area surveyed within the State of
Oregon (D. Goheen and A. Saavedra personal observations).
Ten visually healthy trees were selected for this study (approximately of 20 cm average
diameter at breast height). Trees were inoculated the last week of June 2003. Mycelial
plugs from 7 day-old colonies of isolates 0917-2, 4048 and 4074, and agar plugs from
uninoculated CA plates were used as inoculum. Holes (5 mm dia.) were punched through
the bark to the cambium with a cork borer, the agar plug was inserted and the bark
replaced. A moist piece of cheese cloth was placed over the inoculation site and covered
with aluminum foil secured in place with tape (Brasier and Kirk, 2001; Hansen et al.
2005). The isolates and the control were randomly placed around the bole at cardinal
27
directions and at either 15 or 45 cm from the soil surface. Five weeks after
inoculation tree outer bark was removed using a drawknife to expose the phloem.
Lesions in the inner bark were measured horizontally and vertically from the inoculation
point and the lesion area was then calculated.
Seedling inoculation trial
Due to lack of sources of seeds or seedlings of golden chinquapin for this study, seed
was collected in the wild. During the months of October and November 2003, hundreds
of fruits from several trees and shrubs of golden chinquapin in Oregon and northern
California, were collected (Table 2.2)
Table 2.2 List of trees and shrubs of golden chinquapin and locations were fruits were
collected
Tree ID
Location
Latitude
LA-i
Limi Co., OR
N 44.25376 W 122.36595
Tree
Oct. 1, 2003
LB-i
Siskiyou Co., CA
N 41.958 17
W 123.12238
Tree
Oct. 8, 2003
LB-2
Siskiyou Co., CA
N 4 1.94930
W 123.11497
Tree
Oct. 8, 2003
LC-2
Jackson, Co., OR
N 42.3 1378
W 122.29383
Shrub
Oct. 9, 2003
LD-i
Jackson, Co., OR
N 42.38452
W 122.36751
Tree
Oct. 9,2003
LD-2
Jackson, Co., OR
N 42.38784
W 122.35649
Tree
Oct. 9, 2003
LD-1 (1028)
Jackson, Co., OR
N 42.38452
W 122.36751
Tree
Oct. 28, 2003
Longitude
Tree/shrub
Date collected
Collected fruits were dried and nuts extracted at USDA-Forest Service Dorena Genetic
Resource Center. Nuts that were released from the burrs after the drying process were
checked for damage, mainly caused by insects. Clean seeds were planted in 5 cm dia. x
28
17 cm depth DeepotsTM (HummertTM International Earth City, MO) in the greenhouse
at Dorena.
Only 60 seeds germinated out of several hundred planted. After 8 months, the
surviving seedlings were transported to an Oregon State University greenhouse. At this
facility they were maintained and prepared for the inoculation trials.
Two months before the trial, seedlings were observed to be infected with powdery
mildew. They were sprayed twice with a 1:1 (v/v) solution of Quintec (Dow
AgroSciences), a protective fungicide used for the control of powdery mildew on grapes
and hops (Pscheidt and Ocamb, 2005), and KaligreenTM (Toagosei, Co., LTD, Japan), a
potassium bicarbonate salt that in trials has reduced the infection by powdery mildew in
flowering dogwood (Mmbaga and Sheng, 2002; Mmbaga and Suave, 2004).
Seedlings were inoculated with a zoospore suspension from isolates 4048 and 4074.
To induce sporangia, mycelial plugs from 7-day-old colonies on CA were placed into
sterile 9 centimeter petri dishes and flooded with ten ml clarified V8 broth and incubated
for two days at 18°C (Ito and Kudo, 1994). Each plate was rinsed 3-5 times using
deionized water and then flooded with 10 ml of soil extract water (ig soil / 100 ml
deionized water) and incubated for another two and one half days.
To induce zoospore release, the soil extract was replaced with cold deionized water
(water was chilled in a freezer for
1 hour) and the plates were incubated for 2 hours at
4°C then returned to room temperature. The concentration of released zoospores was
determined using a (Spencer Bright-line) hemacytometer (AO Instrument Co., Scientific
Instrument Division. Buffalo, NY. USA) with the help of a light microscope. The
zoospore concentration was calculated to be
1x104 per
ml.
29
On August
1st,
2005, 40 one-year-old chinquapin seedlings growing in pots were
selected to be inoculated with the two isolates grown for this trial. A hole in the soil of
each pot was opened using a glass rod. Ten ml of zoospore suspension from each isolate
were added to each pot. Ten additional seedlings that served as controls were treated
with 20 ml of one-percent soil extract. All seedlings were immediately flooded with tap
water. After 24 hours the flooding water was drained and the seedlings were watered
twice every day for 19 days and then the pots were flooded again for another 24 hours.
After that plants were watered twice a day again. Seedling mortality was recorded daily
for the duration of the study. During and after the experiment dead seedlings were
examined for rotting roots and inner bark stain. Diseased tissues were plated in CARP,
incubated in the dark at room temperature and observed under the light microscope.
2.3 Results
2.3a. Isolate identification
Colony description
Colony morphology of the chinquapin isolates in this study closely resembled the tester
isolates and the published descriptions of P. cambivora (Waterhouse, 1956; Waterhouse,
1963; Erwin and Ribeiro, 1996). Colonies growing in CA were cottony, with moderate
to profuse aerial mycelium and no particular pattern in the agar (Fig 2.1). The mycelia
were coralloid (Fig 2.2a) with distinct hyphal swellings.
30
Fig 2.1. Colony morphology of isolates of Phytophthora cambivora at room temperature
Mating type testing
Mating type pairing revealed both Al and A2 mating types among the chinquapin
isolates from Oregon forests. The two isolates recovered from the Butte Falls area (Table
2.1) formed reproductive structures when paired against known Al isolates of
Phytophthora cambivora and P. cinnamomi after a little more than a week growing in CA
The resulting oogonia with "warts" and the two celled amphigynous antheridia
morphologically resembled those of P. cambivora (Fig 2.2a, b, c). It was concluded that
isolates 0917-2 and 4048 were mating type A2. Isolate 4074, which was recovered from
31
soil from a stand with healthy golden chinquapin trees approximately 150 miles north
of Butte Falls, did not form oogonia when paired with any of the tester isolates. It did
form sexual structures when paired with the other two chinquapin isolates, 4048 and
0917-2, however. As a result, we concluded that isolate 4074 was mating type Al.
Oogonial diameters were measured for 50 oogonia from each pairing test. Average
oogonial diameter of isolate 09 17-2 was 40.7 tm; oogonial diameters of isolate 4048
averaged 43.3 .tm and isolate 4074 averaged 43.8 tm. All of these measurements are
close to those described by Stamps et al. (1990) and Erwin and Ribeiro (1996) for
Phytophthora cambivora. Table 2.3 summarizes the characters of the reproductive
organs obtained from this study.
Table 2.3 Comparison of the characters of the isolates from this study and those reported
in the literature
Oogonia
Sporangia
CURRENT
1990
1996
STUDY
43-63
41-44
ornamented
ornamented
ornamented
position
amphygynous
amphygynous
amphygynous
No. cells
2
2
2
length (.im)
> 75
40-75
65
morphology
ovate, ellipsoid
ovate, ellipsoid
ovate, ellipsoid
non
non
non
hyphal swellings
present
present
present
growth temperature(°C)
23-27
22-24
20-25
papillate
Colonies
ERWINAND RIBEIRO
40-50
diameter (rim)
morphology
Antheridia
STAMPS ETAL.
32
Fig 2.2 Amphigynous antheridia and oogonia of isolates of Phytophthora cambivora
(aisolate 0917-2; b = isolate 4048; cisolate 4074. Bar = 30 microns)
33
Sporangia description
After two and one half days incubation at 18°C in soil extract water in the dark, isolates
4074 and 4048 formed sporangia in abundance. Isolate 09 17-2 also formed sporangia but
only after 4-5 days incubation and sporangia were not as numerous as the other two
isolates. Sporangia for all three isolates were non-papillate, ovoid in shape, non-cacluceus
and with an average length of 65 gm.
d. Optimal growing temperature
Optimal growing temperature for the chinquapin isolates was 25°C (Table 2.4 and Fig
2.3). At this temperature, isolates grew from 2.2 to 4.4 mm/day on CA. All isolates grew
at 5°C and 35°C, but much more slowly. These growth characteristics are comparable to
published values for P. cambivora (Table 2.3).
34
Table 2.4. Comparison of growth rate among isolates of P. cambivora in this study
Name of
Isolate
0917-2
Temperature
(Celsius)
mm'
mm2
5
0.2
25
0.4
2.2
3.8
4.3
4.4
0.5
0.3
0.8
2.4
4.8
6.2
6.9
30
1.9
35
0.6
10
15
20
25
30
35
4048
5
10
15
20
4074
5
1.3
10
15
3.5
5.8
6.7
8.8
1.1
1.9
2.1
2.2
0.2
0.1
0.4
1.2
2.4
3.1
3.4
0.9
0.3
0.6
1.8
2.9
20
3.3
25
4.4
30
1.0
0.5
35
0.3
0.7
Mean of 2 days growth rate along 4 radii from 3 plates per isolate
2
Mean daily growth rate along 4 radii from 3 plates per isolate
Figure 2.3. Mean daily growth rate (mm) from three plates per 3 isolates of P. cambivora
tested in this study
35
d. ITS rDNA sequencing
The rDNA- ITS sequence of the chinquapin isolates matched from 99% to 100% with
GenBank sequences of Phytophthora cambivora. Isolate 0917-2 and 4048 were 100%
homologous (619 of 619 bps and 627 of 627 bps respectively) with Phytophthora
cambivora isolate AG45 clone 2 (GenBank accession number AY787029). Isolate 4074
was 99% homologous with AG45 clone 2 (783 of 784 bps). For isolates 0917-2 and
4074, nine often best sequence matches from GenBank were for P. cambivora
accessions. The only exception was a sequence for Phytophthora a/ni subsp. mul4formis
(GenBank accession number AY689 136.1). For isolate 4048, besides P. alni subsp
multformis, Phytophthorafragariae was among the top 10 returns. P. fragariae and P.
cambivora are presumably parents of hybrid P. alni. Although the latter also possesses
ornamented oogonia and it is closely related to P. cambivora (Brasier et al. 2004), P. alni
is a homothallic species whereas P. cambivora is a heterothallic species like the
chinquapin-related isolates. Phytophthorafragariae, is closely related to P. cambivora
but, it doesn't have an ornamented oogonium and it is a homothallic species (Stamps et
al. 1990; Cooke et al. 2000).
2.3b. Pathogenicity testing
Tree inoculation trial
Five weeks after inoculations, the bark of inoculated golden chinquapin trees was
removed using a draw knife, revealing areas of necrotic ph!oem around the inoculation
points (Fig 2.4). Lesions were dark brown and diamond shaped. A limited area of
necrotic phloem was observed around control wounds. Control lesion area averaged 1.1
36
cm2. The mean lesion area for the inoculated isolates was 171.4 cm2. Phytophihora
was reisolated on CARP+ from lesions on 8 out of 10 inoculated trees. Recovered
isolates were morphologically similar to colonies of the same isolates growing in the
laboratory.
Measurements of the lesions on the inner bark were collected and lesion area calculated
(Fig 2.5; Table 2.5). Analysis of the effect of the location of the inoculations in trees (15
cm from ground or 45 cm from ground) and the aspect of inoculations (N, S, E, or W)
showed no significance (p-values 0.27 and 0.77 respectively), so the data were pooled for
further analysis. The difference in lesion area between inoculated wounds and control
wounds was significant (p-value = 0.0000 14) (Fig 2.6; Table 2.6). However, there was
no significant difference in lesion area caused by the isolates used in this study (Fig 2.7;
Table 2.7)
Fig 2.4. Bark lesions on bole of golden chinquapin trees: (a) bark lesion caused by P.
cambivora isolate inoculated in bark; (b) control, plain carrot agar in same tree
37
Table 2.5. Bole lesion area (cm2) by tree per isolate of P. cambivora inoculated in the
bark of ten golden chinquapin trees
Tree
identfIcation
Ml
M2
M3
M4
M5
Ti
12
T3
Isolate
identflcation
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
4074
09 17-2
14
15
4048
ctrl
4074
0917-2
4048
ctrl
4074
0917-2
4048
ctrl
Location
in the bole
Lower
Upper
Upper
Lower
Upper
Lower
Upper
Lower
Lower
Upper
Upper
Lower
Upper
Upper
Lower
Lower
Lower
Upper
Upper
Lower
Upper
Upper
Lower
Lower
Upper
Lower
Lower
Upper
Upper
Upper
Lower
Lower
Upper
Lower
Upper
Lower
Lower
Upper
Upper
Lower
Aspect
South
West
East
North
West
North
East
South
East
North
South
West
South
North
West
East
East
South
North
West
South
North
East
West
East
South
North
West
South
North
West
East
West
South
East
North
West
South
North
East
Length
(cm)
35.1
41.8
39.1
0.4
20
22.5
18.7
0
24.8
27
27.1
0
41.4
38.9
28.3
0
26.5
28
30
5.2
37.4
36.2
20.1
1.1
25.5
32.8
29.7
0.4
38.3
20.7
33.7
0.9
35.1
24.8
33.3
0.3
26.1
19.7
31.3
0
Width
(cm)
10.3
10.3
12.9
Lesion'
area (cm2)
0.3
2.2
5.5
0.1
16.3
0
21.5
19.7
25.5
180.8
215.3
252.2
22.0
61.9
152.4
0.0
0
17.7
266.6
266.0
345.5
0.0
366.4
16.1
313.1
16.3
0
7.5
230.6
0.0
99.4
11
154.0
144.0
10.7
203.8
175.6
57.3
9.6
4.1
10.9
9.7
5.7
0.2
9.2
10.5
10.4
0.1
117.3
172.2
154.4
0.2
0.0
7.3
8.7
6.4
0.4
14.4
8.2
9.9
0.2
14.8
139.8
ii.!
2.1
0
Lesion area is the product of length x width divided by 2
90.0
107.8
0.2
252.7
101.7
164.8
0.0
193.1
109.3
32.9
0.0
38
Figure 2.5. Basal lesion area (cm2) by tree per isolate of P. cambivora inoculated in the
bark of ten golden chinquapin trees
400
350
H
300
E
250
J
100 -J
50
0
Ml
M2
M3
M4
M5
TI
T2
T3
T4
T5
Tree No.
Table 2.6. Analysis of variance for isolate effect (including the control as an isolate) on
mean lesion area in inoculated boles of Chrvsolevis chrvsovhvlla
Source
df
P
Ms
F
Isolates
3
11.94495
0.000014
73344.27
Residuals
36
6140.19
Total
39
79484.46
39
Table 2.7. Analysis of variance for isolate effect on mean lesion area on inoculated boles
of Chrysolepis chrysophylla with isolates of P. cambivora (control not included)
P
F
Source
Ms
df
0.86
0.15
Isolates
2
1227.9
Residuals
27
8183.2
Total
29
9411.1
Figure 2.7. Mean lesion area (cm2) of 3 isolates per ten trees
300
200
100
0917-2
4048
Isolate
4074
Analysis of variance showed that there was a significant difference in lesion area
between trees (Fig 2.5; Table 2.8) with a p-value of 0.000933.
Table 2.8. Analysis of variance for tree effect on mean lesion area in boles of Chrysolepis
chrysophylla inoculated with isolates ofF. cambivora (control not included)
Source
df
P
Ms
F
Tree No.
0.000933
9
17488.1
5.30
Residuals
20
3300.4
Total
29
20788.5
Seedling inoculation
100% of seedlings that were soil inoculated with isolates 4048 and 4074 were dead
after 38 days under greenhouse conditions (Fig 2.8; Table 2.9); there was no mortality
among control seedlings that were treated with soil extract water only. First symptoms
were visible after 11 days. Symptoms included yellowing of leaves that later turned
brown and either dropped or remained on stems but dried. The roots were completely
brown and rotted in comparison to the roots of the control seedlings that showed no
necrotic symptoms (Fig 2.9). Observations of the inner bark of roots showed necrosis
caused by the inoculated isolates. Examinations of the inner bark of stems also showed
lesions red to brown in color and girdling the stems. The lesions were determined to be
coming from the main roots of inoculated seedlings.
41
Fig 2.8. One-year-old golden chinquapin seedlings 38 days after inoculation with
isolates of P. cambivora under greenhouse conditions: a) control seedling; b) inoculated
(b)
Fig 2.9. Root conditions of one-year-old golden chinquapin seedlings 38 days after
inoculation with isolates of P. cam hi vora under greenhouse conditions: a) control
g
Table 2.9. Mortality of golden chinquapin seedlings inoculated with isolates of
P. cambivora
Number of seedlings Dead Alive
Inoculated seedlings
40
40
0
Control seedlings
10
0
10
42
2.4 Discussion
The results of this study confirm that the Phytophthora species isolated from Oregon
forest soil and from dying golden chinquapin trees is Phytophthora cambivora (Petri)
Buisman. Morphological characteristics of these isolates agreed with those of P.
cambivora. Colonies were moderate to fast growing with no specific patterns and with
aerial mycelium which gave them a cottony appearance. Mycelium presented numerous
hyphal swellings and no chiamydospores were observed. Oogonia were distinctively
ornamented with a diameter ranging from 40.7 gm to 43.8 im and the two celled
antheridia were amphigynous. Sporangia, which were produced in large numbers, were
non papillate, ellipsoid to ovoid, non deciduous, with an average length of 65tm.
The chinquapin isolates showed some variation in growth rate at different temperatures.
Isolate 4074, which was recovered from soil in the northern range of golden chinquapin,
had a faster growth rate than isolate 0917-2 from Southern Oregon which was also
recovered from soil. All three isolates showed little growth at 5°C and above 30°C. The
optimal growing temperature ranged between 20°C to 25°C as has been reported for this
species (Erwin and Ribeiro, 1996). Both mating types were morphologically similar and
their growth rate was affected in the same maimer depending upon the temperature. It
can't be determined from the isolates used in this study that mating type Al of
Phytophthora cambivora is a faster grower than A2 as has been reported for other species
of Phytophthora (Brasier, 2003). In addition, the ITS rDNA sequencing results
confirmed that the isolates recovered from soil, whether surrounding healthy or declining
golden chinquapin, as well as the isolate recovered from an active lesion margin belong
43
to P. cambivora as they matched from 99% to 100% bp with GenBank sequences of
Phytophthora cambivora (AG45 clone 2).
There have been other reports of Phytophthora cambivora in the Pacific Northwest.
Researchers in Oregon and Washington have associated P. cambivora with root rot and
stem canker in Noble fir in Christmas tree plantations (Hamm and Hansen, 1987;
Chastagner et al. 1995). In addition, Phytophthora cambivora has been recovered from
forest soil around Douglas-fir trees with basal cankers (Hansen pers. comm.). Isolates
used in the present study were from forest settings associated with this new canker
disease on golden chinquapin as well as from forest soil around apparently healthy
chinquapins in an area where unusual mortality has not been noted.
Both mating types of P. cambivora have been reported from the Pacific Northwest.
Mating type Al of P. cambivora was frequently recovered from dead roots and stem
cankers in noble fir Christmas tree plantations (Hansen pers. comm.). In addition, an
isolate of mating type A2 from noble fir in Oregon (ATTC 46719) was cited by
Oudemans and Coffey (1991) in their isozyme comparison study, and in the present
study, both mating types of P. cambivora were recovered from Oregon forest soil in
geographically distant locations. Further surveys across the range of golden chinquapin
will be needed to verify the distribution of P. cambivora mating types.
Results of tree and seedling inoculations support the original hypothesis that golden
chinquapin is susceptible to Phytophthora cambivora. The method used for inoculating
seedlings, adding zoospore suspension directly into the soil, gave results comparable to
other pathogenicity studies with P. cambivora (Jung et al. 2003; Jung et al. 2005). In
these studies, Phytophthora cambivora was incorporated into the soil and caused root rot
44
and mortality (85-100% and 70-90 to 100% respectively) on seedlings of European
beech (Fagus sylvatica L.). The current study showed that seedlings of golden
chinquapin were also highly susceptible (100% mortality) to P. cambivora under
greenhouse conditions. Examinations of roots showed that they were highly rotted and
their biomass significantly reduced (Fig 2.9) compared to control roots. In addition,
extensive necrotic lesions were also found in the inner bark of every killed seedling.
Because isolates were mixed for the seedling inoculations, differences in aggressiveness
of the isolates ofF. cambivora could not be addressed in this study. Recovery of P.
cambivora from inoculated seedlings successfully completed an objective of this study by
satisfying Koch's postulates. However, it is also important to note that the seedlings used
in this study represented a very limited portion of the possible variation of responses that
a larger population of golden chinquapin seedlings might have provided. Due to the lack
of availability of seedlings from commercial nurseries and the low rate of seed
germination, only 50 seedlings from four specific areas that were readily accessible by
roads were used in this study. A more detailed study representing a more diverse
population from the entire range of golden chinquapin is necessary to answer more
questions regarding the level of susceptibility of golden chinquapin to P. cambivora.
A short study during the summer of 2002 (Saavedra and Goheen; no data included in
this work) collected data regarding the behavior of this disease in the area in which P.
cambivora is causing the most damage. This study suggested a relationship between road
proximity and incidence of the disease. Several surveys conducted during the following
two summers within the range of golden chinquapin also suggested that the disease might
be related to road proximity.
45
Turchetti (1988) reported that P. cambivora penetrated chestnuts through the fine
roots. The results of drenching zoospores directly into the soil suggest that penetration of
this pathogen into seedlings of golden chinquapin is also through fine roots. Observations
of the lesions on stems in the forest supported this possibility, since the infection on the
stem was always traceable to infected main roots. The mechanism of penetration of
encysted zoospores into the fine roots of golden chinquapin cannot be explained in this
current work but a study described by Casares et al. (1994) reported that hyphae of P.
cambivora grew intercellularly in callus cells of European chestnut (Castanea sativa
Miller) by the action of degrading enzymes produced at the tip of advancing hypbae and
also intracellular infection of callous cells via haustorial formation. This study could
provide insight to explain modes of infection of P. cambivora into the roots of golden
chinquapin seedlings.
Field inoculation trial results also confirmed the hypothesis that golden chinquapin is
susceptible to P. cambivora. Lesions were observed in the inner bark of inoculated trees
P. cambivora was recovered from the margins of active lesions. Although the boles of
inoculated trees were almost girdled by the pathogen by the end of the trial period, the
trees were still alive. Crown symptoms were observed, however, including leaf chlorosis
and some branch flagging. It is believed that the infection caused by this pathogen would
have led to the eventual death of inoculated trees. Two years after the end of this trial the
inoculated trees were dead (pers. observations) but the mechanical girdling of trees
during lesion measurement undoubtedly contributed to the rapid mortality.
46
There are questions that need to be addressed in further studies for achieving a better
understanding of this new disease. Is the apparent correlation between road proximity
and tree mortality real? If it is real, is it due to predisposition, or transport of the
pathogen along the roads? Another question that remains unanswered is the
aggressiveness of this disease. There is little information on how quickly mature trees
might be killed after initial infection by P. cambivora. Results of this study showed rapid
advance of Phytophthora cambivora through phloem tissues suggesting that girdling of
trees could occur within months of bark infection but does not answer the question of
how long it takes for the pathogen to move out of the roots, or how long it takes girdled
trees to die.
In summary, Phytophthora cambivora is present in Oregon forest soil and both mating
types are represented. Both inoculation methods used, bark inoculation of mature living
trees or root infection of seedlings growing in soil infested with a zoospore suspension,
resulted in the formation of necrotic tissues and in the case of the greenhouse seedling
experiment, 100% mortality. In both instances, the pathogen was successfully recovered
from the infected tissue satisfying Koch's postulates and demonstrating that
Phytophthora cambivora is the causal agent of this new root rot and basal canker disease
of golden chinquapin.
However, further studies need to be conducted for better understanding of this disease
and its ecological impact in Oregon forests.
47
Literature Cited:
Altschul, S.F., Maden, T.L., Schäffer, A.A., Zhang, J., Zhang, A., Miller, W. and
Lipman, D.J. (1997). Gapped-BLAST and PSI-BLAST: a new generation of
protein database search program. Nucleic Acids Res. 25: 33 89-3402.
Anagnostakis, S. (1987). Chesnut blight: the classical problem of an introduced pathogen.
Mycologia 79(1): 23-37.
Barnard, E. L. (2000). Chesnut Blight of Chinkapin in Florida., Fla. Dept. Agric. &
Consumer Serv. Division of Plant Industry. Plant Pathology Circular No. 402.
Brasier, C.M. (2003). Sudden oak death: Phytophthora ramorum exhibits transatlantic
differences. Mycol. Res. 107(3): 258-259.
Brasier, C. M. and Kirk, S. (2001). Comparative aggressiveness of standard and variant
hybrid alder Phytophthoras, Phytophthora cambivora and other Phytophthora
species on bark of Alnus, Quercus and woody hosts. Plant Pathology 50: 218-229.
Brasier, C. M. and Kirk, S. (2004). Production of gametangia by Phytophthora ramorum
in vitro. Mycol. Res. 108(7): 823-827.
Brasier, C. M. Kirk, S., Delcan, J., Cooke, D., Jung, T. and Man In't Veld, W.A. (2004).
Phytophthora alni sp. nov. and its variants: designation of emerging heteroploid
hybrid pathogens spreading on Alnus trees. Mycol. Res. 108(10): 1172 - 1184.
Brockman, C. F. (1958). Golden chinquapin. University of Washington Arboretum
Bulletin 21: 46-47, 70.
Casares, A., Gonzalez, A. and Sanchez Tames, R. (1994). Ultrastructure of calli from
resistant and susceptible chestnut inoculated with Phytophthora cambivora. J.
Phytopathology 142: 19-26.
Chastagner, G., Hamm, P. and Riley, K. (1995). Symptoms and Phytophthora spp.
associated wit root rot and stem canker of Noble fir Christmas trees in the Pacific
Northwest. Plant Disease 79(3): 290-293.
Cooke, D.E.L., Drenth, A., Duncan, J.M., Wagels, G. and Brasier, C.M. (2000). A
molecular phylogeny of Phytophthora and related oomycetes. Fung. Gen. Biol.
30: 17-32.
Crandall, B., Gravatt, G.F. and Ryan, M.M. (1945). Root disease of Castanea species and
some coniferous and broadleaf nursery stocks, caused by Phytophthora
cinnamomi. Phytopathology 35: 162-180.
De Cock, A. W. A. M. and Levesque, C.A. (2004). New species of Pythium and
Phytophthora. Studies in Mycology 50: 48 1-487.
Erwin, D. and Ribeiro, 0. (1996). Phytophthora Diseases Worldwide. St. Paul,
Mirmesota. USA., APS Press. 562p.
Fan, D., Bills, G., Chamuris, G. and Rossman, A. (1989). Fungi on Plants and Plant
Products in the United States., The American Phytopathological Society. St. Paul,
MN. USA. 1252 p.
Hamm, P.B. and Hansen E.M. (1987). Identification of Phytophthora spp. known to
attack conifers in the Pacific Northwest. Northwest Science 61(2): 103-109.
Hansen, E. M. (2001). Phytophthora in the Americas. In: Phytophthora in Forests and
Natural Ecosystems.
International IUFRO Working Party 7.02.09 Meeting.
Albany, W. Australia 30th Sept. -5 Oct 2001. McComb, J., Hardy, G. and
Tommerup, I. (eds.) (Murdock University Print) Pp: 19 - 24.
Hansen, E. M., Parke, J.L. and Sutton, W. (2005). Susceptibility of Oregon forest trees
and shrubs to Phytophthora ramorum: A comparison of artificial inoculation and
natural infection. Plant Disease 89: 63-70.
Hubbard, R. L. (1974). Castanopsis (D.Don) Spach, chinkapin. In: Seeds of Woody
Plants in the United States. C.S. Schopmeyer tech. coord. Washington D.C.,
USDA, Agriculture Handbook 450: 276-277.
Ito, M. and Kudo, A. (1994). Promotion of zoosporangium formation in Phytophthora
cambivora (Petri) Buisman, pathogen of apple crown rot in Japan. Bull. Fruit Tree
Res. Stn. 26: 71-82.
Jung, T., Cook, D.E.L., Blaschke, H., Duncan, J.M. and Owald, W. (1999).
Phytophthora quercina sp. nov., causing root rot of European oaks. Mycol. Res.
103(7): 785-798.
Jung, T., Hansen, E.M., Winton, L., Owald, W. and Delatour, C. (2002). Three new
species of Phytophthora from European oak forest. Mycol. Res. 106(4): 397-411.
Jung, T., Nechwatl, J., Cooke, D.E., Hartmann, 0., Blaschke, H., OI3wald, W.F., Duncan,
J.M. and Delatour, C. (2003). Phytophthora pseudosyringae sp. nov., a new
species causing root and collar rot of deciduos tree species in Europe. Mycol. Res.
107(7): 772-789.
Jung, T., Hudler, G., Jensen-Tracy, S.L., Griffiths, H.M., Fleiscbman, F. and Osswald,
W. (2005). Involvement of Phytophthora species in the decline of European
beech in Europe and the USA. Mycologist 19(4): 159-166.
49
Kaosiri, T., Zentmyer, G.A. and Erwin, D.C. (1978). Stalk length as a taxonomic
criterion for Phytophthorapalmivora isolates from cacao. Can. J. Bot. 56: 17301738.
Kellam, M.K. and Zentmyer, G.A. (1986). Comparisons of single-oospore isolates of
Phytophthora species from naturally infected cocoa pods in Brazil. Mycologia 78
(3): 351-358.
Kruckeberg, A. R. (1980). Golden Chinquapin (Chrysolepis chrysophylla) in Washington
State: A species at the northern limit of its range. Northwest Science 54 (1): 4-16.
Mc Kee, A. (1990). Castanopsis chrysophylla (Dougl-) A.DC. Giant Chinkapin. j
Silvics of North America, Burns, R. and Honkala, B. (eds). USDA-Agricultural
Handbook No.654. 8'77p.
Mmabaga, M. and Sheng, H. (2002). Evaluation of biorational products for powdery
mildew management in Cornusfiorida. J. Environ. Hort. 20(2): 113-117.
Mmabaga, M. and Suave, R.J. (2004). Management of powdery mildew in flowering
dogwood in the field with biorational and conventional fungicides. Can. J. Plant
Sci. 84: 837-844.
Newhook, F. J., Waterhouse, G.M. and Stamps, D.J. (1978). Tabular key to the species of
Phytophthora De Bary. CMI. Mycological Papers No. 143.
Oudemans, P. and Coffey, M.D. (1991). Isozyme comparison within and among
worldwide sources of three morphologically distinct species of Phytophthora.
Mycol. Res. 95(1): 19-30.
Payne, J., Johnson, G. and Miller, G. (1993). Chinkapin: potential new crop for the south.
In: New Crops. Janick and Simon (eds). Wiley, NY: 500-505.
Pirone, P. (1940). Phytophthora disease of maple. Phytopathology 30: 19.
Pscheidt, J. W. and Ocamb, C. (2005). Pacific Northwest Plant Disease Management
Handbook. Corvallis, OR. Extension Service of Oregon State University. 537p.
Russo, D. (1979). Plant Galls of the California Region. Pacific Grove, CA, The Boxwood
Press. 2O3p.
Santini, A., Barzanti, G.P. and Capretti, P. (2001). A new Phytophthora root disease of
alder in Italy. Plant Disease 85(5): 560.
Scheffer, T. H. (1961). Search for the chinquapin in the State of Washington. University
of Washington Arboretum Bulletin 24: 88, 96-97.
50
Staden, R. (1996). The Staden sequence analysis package. Mol. Biotechnol. 5: 233241.
Stamps, D. J., Waterhouse, G.M., Newhook, F.J. and Hall, G.S. (1990). Revised Tabular
Key to Species of Phytophthora. Kew, UK: CMI. Mycological Papers. 28p.
Turchetti, T. (1988). Phytophthora cambivora (Petri) Buisman. In: European Handbook
of Plant Diseases. Smith, I.M., Dunez, J., Phillips, D.H. Lelliot, R.A. and Archer,
S.A. (eds). Blackwell Scientific Publications. 2' edition. 2l5p.
Vettraino, A. M., Vannini, A., Anselmi, N. and Fuibright, D.W (2000). Survey of
Phytophthora species from soil surrounding diseased chesnut species in North
America. In: Phytophthora Diseases of Forest Trees. 1st International IUFRO
Working Party 7.02.09 Meeting. Corvallis, Oregon: Oregon State University.
USA. Hansen, E.M. and Sutton, W. (eds). Pp: 145-147.
Waterhouse, G.M. (1956). The genus Phytophthora diagnoses (or descriptions) and
figures from the original papers. C.M.I. Mycological papers No. 12. l2Op.
Waterhouse, G. M. (1963). Key to the species of Phytophthora de Bary. Mycol. Pap. No.
92: 22p.
Werres, S., Marwitz, R., Man In't Veld, W., De Cock, A.W.A.M., Bonants, P., De
Weerdt, M., Themann, K., Ilieva, E. and Baayen, R. (2001). Phytophthora
ramorum sp. nov., a new pathogen on Rhododendron and Viburnum. Mycol. Res.
105(10): 1115-1165.
White, T. J., Bruns, T., Lee, S. and Taylor, J. (1990). Amplification and direct sequencing
of fungal ribosomal RNA genes for phylogenetics. In: PCR Protocols: a Guide to
Methods and Applications. hmis, M.A., Geldfand, D.H., Sninsky, J.J., and White,
T.J. (eds). San Diego: Academic Press: 3 15-321.
Winton, L. M. and Hansen, E.M. (2001). Molecular diagnosis of Phytophthora lateralis
in trees, water, and foliage baits using multiplex polymerase chain reaction.
Path. 31: 275-283.
Zentmyer, G. A., Leary, J.V., Kiure, L.J. and Grantham, G.L. (1976). Variability in
growth of Phytophthora cinnamomi in relation to temperature. Phytopathology
66: 982-986.
51
Summary and Conclusion
Chapter 3
Phytophthora cambivora has been known as a pathogenic species on trees of economic
importance, especially European chestnut, since it was first described by Petri in 1917
(Cited in Tucker, 1933). Later other authors reported this pathogen affecting other tree
species of economic and ecological importance (Pirone, 1940; Mircetich et al. 1974;
Jeffers and Aldwinckle, 1988; Chastagner et al. 1995; Sbafizadeh and Kavanagh, 2005).
In all of these reports P. cambivora was causing root rot and basal canker to its hosts.
Golden chinquapin, a native tree of the western United States, is not known to be
affected by many pests other than a few heart-rotting fungi and a few insect pests
(Hepting, 1971; McKee, 1990; Sinclair and Lyon, 2005). This species is very resistant to
chestnut blight, caused by Cryphonectria parasitica, in contrast to its close relatives,
Castanea species.
However, in recent years serious pockets of mortality affecting golden chinquapin trees
have been observed (D. Goheen pers. comm.). Infected trees showed crown symptoms
including leaves turning from green to bright red before being dropped, or branch
flagging, with the dead leaves retained. Another symptom was the presence of necrotic
tissues in the inner bark of stems and roots.
Initial isolations from active growing cankers and from surrounding soil yielded a
Phytophthora species morphologically resembling P. cambivora. This current study was
52
undertaken to establish the true identity of the recovered fungus, and to determine its
pathogenicity on golden chinquapin by satisfying Koch's postulates.
Several approaches were pursued to identify the isolates. Three chinquapin isolates
were characterized and compared to published descriptions. Two of the isolates were
recovered from the epidemic area (Butte Falls, OR), one from soil associated with dead
trees and the other from necrotic inner bark of an infected golden chinquapin. They were
labeled as isolates 0917-2 and 4048 respectively. A third isolate was recovered from soil
near healthy-appearing chinquapin trees in the northern range of the tree (Linn Co. OR).
This isolate was identified as 4074.
The isolates were grown in carrot agar and their morphological characteristics were
compared with those of known isolates of Phytophthora cambivora. The chinquapin
isolates produced aerial mycelium giving the colonies a cottony appearance, and the
hyphae often exhibited coralloid hyphal swellings. These characters were also present on
the known isolates of P. cambivora and matched those described in the literature
(Waterhouse, 1956; Stamps et al. 1990).
To induce sporangia, a technique modified from Ito and Kudo (1994) was used.
Numerous sporangia were formed. Sporangia were ellipsoid, non papillate, non
caducous, with simple sporangiophores and measured in average 65tm in length.
The isolates' mating types were also resolved by pairing against tester isolates of
known mating type. As a result, isolates 09 17-2 and 4048 were determined to be of
mating type A2 whereas isolate 4074 was determined as mating type Al. The resulting
oogonia were ornamented as described for this species (Ho et al. 1977; Stamps et al.
1990) with an average diameter that ranged from 40.7 J.tm to 43.8p.m. The antheridia
53
were two celled and amphigynous, again matching published descriptions for
Phytophthora cambivora.
Growth rate testing results indicated that the optimal growing temperature for the
chinquapin isolates ranged from 20° 25°C, and that growth was restricted or stopped at
extreme temperatures of 5°C and above 3 0°C. These temperature values agreed with the
growing temperatures recorded for this pathogenic species (Erwin and Ribeiro, 1996).
Another approach used for establishing the identity of the chinquapin isolates was
sequencing their ITS r-DNA, as used by other researchers for identifying suspected or
unknown species (Winton and Hansen, 2001; Hansen et al. 2003; De Cock and Lévesque
2004; Brasier et al. 2005; Greslebin et al. 2005). All three isolates showed high
homology with Phytophthora cambivora isolate AG45 clone 2 (GenBank accession
number AY787029). The molecular analysis in combination with morphological and
physiological observations supported the conclusion that all three of the isolates, from
dying chinquapin trees and associated soil, and from soil around healthy trees, belonged
to Phytophthora cambivora.
Pathogenicity tests confirmed that Phytophthora cambivora was the casual agent of this
new canker disease. All 40 seedlings of golden chinquapin inoculated with zoospore
suspensions of Phytophthora cambivora died within 38 days. In contrast, all 10 control
seedlings that were treated with a soil extract solution remained healthy. P. cambivora
was recovered from rotting roots and stems of the inoculated seedlings but not from
control seedlings, completing Koch's postulates.
Inoculation of living, mature chinquapin trees in the field also demonstrated that P.
cambivora is the causal agent of canker disease on golden chinquapin. All 10 golden
54
chinquapin trees inoculated with plugs containing growing mycelia of P. cambivora
developed cankers within weeks after inoculation. Cankers resembled those seen on
dying trees in nature. P. cambivora was isolated from lesions on inoculated trees but not
from control wounds, satisfying Koch's rules.
In summary, P. cambivora has been isolated, identified and confirmed responsible for a
canker disease on golden chinquapin in Oregon. This is believed to be the first report of
P. cambivora causing disease on golden chinquapin.
Future sug2estions
For achieving a further understanding of the behavior of this disease, it is suggested:
1) To conduct a more extensive study of the relationship between road proximity and
severity of the disease in stands where mortality is evident.
2) To make an extensive survey throughout the natural range of golden chinquapin to
determine the impact that this disease may have already caused in nature.
3) To carry out more inoculation trials that include a better representation of golden
chinquapin populations for identifying resistant individuals and studying possible
mechanisms involved in resistance.
4) To conduct a more detailed soil survey for determining the distribution of
Phytophthora cambivora in Oregon forests.
5) To evaluate the ecological impact that this disease may cause in Oregon forest
communities.
55
Bibliography
Altschul, S.F., Maden, T.L., Schaffer, A.A., Zhang, J., Zhang, A., Miller, W. and
Lipman, D.J. (1997). Gapped-BLAST and PSI-BLAST: a new generation of
protein database search program. Nucleic Acids Res. 25: 3389-3402.
Anagnostakis, S. (1987). Chesnut blight: the classical problem of an introduced pathogen.
Mycologia 79(1): 23-37.
Ash, J. (1988). Nothofagus (Fagaceae) forest on Mt. Giluwe, New Guinea. New Zealand
Journal of Botany 26: 245 258.
Barnard, B. L. (2000). Chesnut Blight of Chinkapin in Florida., Fla. Dept. Agric. &
Consumer Serv. Division of Plant Industry. Plant Pathology Circular No. 402.
Betlejewki, F., Casavan, K., Dawson, A., Goheen, D., Mastrofini, K., Rose, D. and
White, D. (2003). A range-wide Assesment of Port-Orford-cedar (Chamaecyparis
lawsoniana) on Federal Lands., Bureau of Land Management, USDA- Forest
Services. l82p.
Bourbos, V. and Metzidakis, I. (2000). Biological control of Phytophthora cambivora
(Petri) Buisman in chestnut tree (Castanea sativa Mill). Ecologia Mediterranea 26
(1-2): 123-127.
Brasier, C. M. (2000). The rise of the hybrid fungi. Nature 405: 134-135.
Brasier, C. M. (2003). Sudden oak death: Phytophthora ramorum exhibits transatlantic
differences. Mycol. Res. 107(3): 25 8-259.
Brasier, C. M. and Jung, T. (2001). Progress in understanding Phytophthora diseases of
trees in Europe. In: Phytoyhthora in Forests and Natural Ecosystems. 2nd
International IIJFRO Working Party 7.02.09 Meeting. Albany. W. Australia 30th
Sept. -5 Oct 2001. McComb, J.. Hardy, G. and Tommerup, I. (eds.) (Murdock
University Print) Pp: 4-18.
Brasier, C.M. and Kirk, 5. (2001). Comparative aggressiveness of standard and variant
hybrid alder Phytophthoras, Phytophthora cambivora and other Phytophthora
species on bark of Alnus, Quercus and woody hosts. Plant Pathology 50: 218-229.
Brasier, C. M. and Kirk, 5. (2004). Production of gametangia by Phytophthora ramorum
in vitro. Mycol. Res. 108 (7): 823-827.
56
Brasier, C. M., Kirk, S., Delcan, J., Cooke, D., Jung, T. and Man In't Veld, W. A.
(2004). Phytophthora alni sp. nov. and its variants: designation of emerging
heteroploid hybrid pathogens spreading on Alnus trees. Mvcol. Res. 108(10):
1172- 1184.
Brasier, C. M., Beales, P. A., Kirk, S. A., Denman, S. and Rose, J. (2005). Phytophthora
kernoviae sp. nov., and invasive pathogen causing bleeding stem lesions on forest
trees and foliar necrosis of ornamentals in the UK. Mycol. Res. 109(8): 853-859.
Brockman, C. F. (1958). Golden chinquapin. University of Washington Arboretum
Bulletin 21: 46-47, 70.
Buisman, C. J. (1927). Root rot caused by Phycomycetes, University of Utrecht. Meded.
Phytopathol. Lab. Willie Commenlin Scholten 11:7. (Rev. Appi. Mycol. 1927,
6:380): 5lp.
Casares, A., Gonzalez, A. and Sanchez Tames, R. (1994). Ultrastructure of calli from
resistant and susceptible chestnut inoculated with Phytophthora cambivora. L
Phytopathology 142: 19-26.
Cavalier-Smith, T. (1986). The Kingdom Chromista: Origin and systematics. In: Progress
in Phycological Research. Round and Chapman (Eds.). Bristol, UK, Biopress.
Vol. 4: 309 - 347.
Chastagner, G., Hamm, P. and Byther, R. (1990a). Symptomology of Phytophthora root
rot and stem canker disease of Noble fir in the Pacific Northewest.
Phytopathology 80(9): 887.
Chastagner, G., Riley, K. and Hamm, P. (1990b). Susceptibility of Abies spp. to seven
Phytophthora spp. Phytopathology 80(9): 887.
Chastagner, G., Hamm, P. and Riley, K. (1995). Symptoms and Phytophthora spp.
associated with root rot and stem canker of Noble fir Christmas trees in the
Pacific Northwest. Plant Disease 79(3): 290-293.
Cooke, D. E. L., Drenth, A., Duncan, J. M., Wagels, G. and Brasier, C. M. (2000). A
molecular phylogeny of Phytophthora and related oomycetes. Fung. Gen. Biol.
30: Pp: 17 -32.
Crandall, B., Gravatt, G. F. and Ryan, M. M. (1945). Root disease of Castanea species
and some coniferous and broadleaf nursery stocks, caused by Phytophthora
cinnamomi. Phytopathology 35: 162-180.
De Cock, A. W. A. M. and Lévesque, C. A. (2004). New species of Pythium and
Phytophthora. Studies in Mycology 50: 48 1-487.
57
Delatour, C. (2001). Phytophthoras and oaks in Europe. In: Proceedings of the 49th
Annual Western International Forest Disease Work Conference. September 1014, 2001. Carmel Mission Inn, Carmel, CA. Pp:16 -30.
Erwin, D. and Ribeiro, 0. (1996). Phytophthora Diseases Worldwide. St. Paul,
Minnesota. USA., APS Press. 562p.
Fan, D., Bills, G., Chamuris, G. and Rossman, A. (1989). Fungi on Plants and Plant
Products in the United States., The American Phytopathological Society. St. Paul,
MN. USA. 1252 p.
Gerrettson-Cornell, L. (1978). Notes on the morphology of an isolate of Phytophthora
cambivora (Petri) Buis. from Australia. Phyton 36(1): 85 - 89.
Greslebin, A. G., Hansen, E. M., Winton, L. M. and Rajchenberg, M. (2005).
Phytophthora species from declining Austrocedrus chilensis forest in Patagonia,
Argentina. Mycologia 97(1): Pp: 218 - 228.
Hamm, P. B. and Hansen, B. (1987). Identification of Phytophthora spp. known to attack
conifers in the Pacific Northwest. Northwest Science 61(2): 103-109.
Hansen, E. M. (2001). Phytophthora in the Americas. In: Phvtophthora in Forests and
Natural Ecosystems. 2nd International IIJFRO Working Party 7.02.09 Meeting.
Albany, W. Australia 30th Sept. -5 Oct 2001. McComb, J., Hardy, G. and
Tommerup, I. (eds.) (Murdock University Print) Pp: 19 - 24.
Hansen, B. M. and Sutton, W. (2002). Log inoculations to asses tree susceptibility to
sudden oak death. Phytopathology 92 (6 Supplement): S33.
Hansen, E. M., Reeser, P. W., Davison, J. M., Garbelotto, M., Ivors, K., Douhan, L. and
Rizzo, D. M. (2003). Phytophthora nemorosa, a new species causing cankers and
leaf blight of forest trees in California and Oregon, USA. Mycotaxon 88: 129138.
Hansen, E. M., Parke, J. L. and Sutton, W. (2005). Susceptibility of Oregon forest trees
and shrubs to Phytophthora ramorum: A comparison of artificial inoculation and
natural infection. Plant Disease 89: 63-70.
Hepting, G. (1971). Diseases of Forest and Shade Trees of the United States. USDA
Agriculture Handbook No. 386. 658p.
Hjelmqvist, H. (1948). Studies on the floral morphology and phylogeny of the
Amentiferae. Bot. Not. Suppl. 2(1): l'7lp.
Ho, H. H., Zentmyer, G. A. and Erwin, D. C. (1977). Morphology of sex organs of
Phytophthora cambivora. Mycologia 69: 641-646.
Hubbard, R. L. (1974). Castanopsis (D.Don) Spach, chinkapin. In: Seeds of Woody
Plants in the United States. C.S. Schopmeyer tech coord. Washington D.C.,
USDA, Agriculture Handbook 450: 276-277.
Ito, M. and Kudo, A. (1994). Promotion of zoosporangium formation in Phytophthora
cambivora (Petri) Buisman, pathogen of apple crown rot in Japan. Bull. Fruit Tree
Res. Stn. 26: 71-82.
Jeffers, S. N. and Aldwinckle, H. 5. (1988). Phytophthora crown rot of apple trees:
Sources of Phytophthora cactorum and P. cambivora as primary inoculum.
Phytopathology 78: Pp: 328 - 325.
Jensen, E. and Ross, C. (1995). Trees to Know in Oregon, Extension Circular 1450.
Oregon State University Extension Services and Oregon Department of Forestry.
l28p.
Jönsson, U., Lunderberg, L., Sonesson, K. and Jung, T. (2003). First record of soilbome
Phytophthora species in Swedish oak forest. For. Path. 33: 175-179.
Jung, T., Cook, D. E. L., Blaschke, H., Duncan, J. M. and Owald, W. (1999).
Phytophthora quercina sp. nov., causing root rot of European oaks. Mycol. Res.
103(7): 785-798.
Jung, T., Blascbke, H. and O3wald, W. (2000). Involvement of soilborne Phytophthora
species in Central European oak decline and the effect of site factors on the
disease. Plant Pathology 49: 706-7 18.
Jung, T., Hansen, E. M., Winton, L., OI3wald, W. and Delatour, C. (2002). Three new
species of Phytophthora from European oak forest. Mycol. Res. 106(4): 397-411.
Jung, T., Nechwatl, J., Cooke, D. E., Hartmann, G., Blaschke, H., Of3wald, W. F.,
Duncan, J. M. and Delatour, C. (2003). Phytophthora pseudosyringae sp. nov., a
new species causing root and collar rot of deciduos tree species in Europe. Mycol.
Re 107(7): 772 - 789.
Jung, T., Hudler, G., Jansen-Tracy, S. L., Griffiths, H. M., Fleischman, F. and Osswald,
W. (2005). Involvement of Phytophthora species in the decline of european beech
in Europe and the USA. Mycologist 19(4): 159-166.
Kaosiri, T., Zentmyer, G.A. and Erwin, D.C. (1978). "Stalk length as a taxonomic
criterion for Phytophthorapalmivora isolates from cacao." Can. J. Bot. 56: 17301738.
59
Kellam, M.K. and Zentmyer, G.A. (1986). Comparisons of single-oospore isolates of
Phytophthora species from naturally infected cocoa pods in Brazil. Mycologia 78
(3): 35 1-358.
Kendrick, B. (2000). The Fifth Kingdom. Newburyport, MA. USA, Focus Publishing. R.
Pullins Co. Y73p.
Kruckeberg, A. R. (1980). Golden Chinquapin (Chrysolepis chrysophylla) in Washington
State: A species at the Northern Limit of its Range. Northwest Science 54(1): 416.
Li, R., Chen, Z., Lu, A., Soltis, D., Soltis, P. and Manos, P. (2004). Phylogenetic
relationship in Fagales based on DNA sequences from three genomes. mt. J. Plant
Sci.165(2): 311-324.
Manos, P., Zhan, Z. and Cannon, C. (2001). Systematics of Fagaceae: phylogenetics test
of reproductive trait evolution. Int. J. Plant Sci. 162(6): 1361-1379.
May, K. J., Drenth, A. and Irwin, J. A. G. (2003). Interspecific hybrids between the
homothallic Phytophthora sojae and Phytophthora vignae. Australasian Plant
Pathology 32: 353-359.
Mc Kee, A. (1990). Castanopsis chrysophylla (Dougi- A.DC. Giant Chinkapin. j
Silvics of North America, Burns, R. and Honkala, B. (eds). USDA-Agricultural
Handbook No.654. 8'77p.
Mircetich, S. M. and Matheron, M. E. (1976). Phytophthora root rot and crown rot of
cherry trees. Phytopathology 66: 549-55 8.
Mircetich, S. M., Matheron, M. E. and Schreader, W. R. (1974). Sweet cherry root rot
and trunk canker caused by Phytophthora spp. Proc. Am. Phytopathol. Soc.
(abstr.) 1: 58.
Mmabaga, M. and Sheng, H. (2002). Evaluation of biorational products for powdery
mildew management in Cornusfiorida. J. Environ. Hort. 20(2): 113-117.
Mmabaga, M. and Suave, R. J. (2004). Management of powdery mildew in flowering
dogwood in the field with biorational and conventional fungicides. Can. J. Plant
Sd. 84: 83 7-844.
Newhook, F. J., Waterhouse, G. M. and Stamps, D. J. (1978). Tabular Key to the species
of Phytophthora De Bary. CMI. Mycological Papers No. 143.
Oudemans, P. and Coffey, M. D. (1991). Isozyme comparison within and among
worldwide sources of three morphologically distinct species of Phytophthora.
Mycol. Res. 95(1): 19-30.
Payne, J., Johnson, G. and Miller, G. (1993). Chinkapin: potential new crop for the south.
In: New Crops. Janick and Simon (eds). Wiley, NY: 500-505.
Petri, L. (1917). Ricerche sulla morfologia e biologia della Blepharospora cambivora,
parassita del Castagno. Atti. R. Accad. Lencei. Rend. Cl. Sci. Fis. Mat. Nat. Ser.
5 26: 297 -299.
Pirone, P. (1940). Phytophthora disease of maple. Phvtopathology 30: 19.
Pscheidt, J. W. and Ocamb, C. (2005). Pacific Northwest Plant Disease Management
Handbook. Corvallis, OR. Extension Service of Oregon State University. 537pp.
Rizzo, D. M., Garbelotto, M., Davidson, J. M., Slaugther, G. W. and Koike, S. T. (2002).
Phytophthora ramorum as the cause of extensive mortality of Quercus spp. and
Lithocarpus densiflorus in California. Plant Disease 86(3): 205 - 214.
Russo, D. (1979). Plant Galls of the California Region. Pacific Grove, CA, The Boxwood
Press. 2O3p.
Santini, A., Barzanti, G. P. and Capretti, P. (2001). A new Phytophthora root disease of
alder in Italy. Plant Disease 85(5): 560.
Scheffer, T. H. (1961). Search for the chinquapin in the State of Washington. University
of Washington Arboretum Bulletin 24: 88, 96-97.
Shafizadeh, S. and Kavanagh, J. A. (2005). Pathogenicity of Phytophthora species and
Pythium undulatum isolated from Abies procera Christmas trees in Ireland. E
Path. 35: 444-450.
Sinclair, W. A. and Lyon, H. (2005). Diseases of Trees and Shrubs. Ithaca and London,
Comstock Publishing Associates, a division of Cornell University Press. 660p.
Smith, J. P. (1977). Vascular Plant Families. Eureka, CA, Mad River Press, Inc. 320p.
Staden, R. (1996). The Staden sequence analysis package. Mol. Biotechnol. 5: 233-241.
Stamps, D. J., Waterhouse, G. M., Newhook, F. J. and Hall, G. S. (1990). Revised
Tabular Key to Species of Phytophthora. Kew. UK: CMI. Mycological papers
No. 162: 1-28.
Tantier, F., O'Brien, J. G., Hernandez, A., Orozco, F. and Rebolledo, 0. (2000).
Phytophthora cinnamomi as a cause of oak mortality in the state of Colima,
Mexico. Plant Disease 84: 394 -398.
Tucker, C. M. (1933). Distribution of the Genus Phytophthora. Univ. MO. Agric. Exp.
Stn. Res. Bull. 184.: 1-80.
Turchetti, T. (1988). Phytophthora cambivora (Petri) Buisman. European Handbook of
Plant Diseases. Smith, I.M., Dunez, J., Phillips, D.H., Lelliott, R.A., and Archer,
S.A. (eds.), Blackwell Scientific Publications. 2nd edition. 215p.
Uphof, J.C.Th. (1959). Dictionary of Economic Plants. Weinheim, H.R. Engelmann
(J.Crammer); New York, Hafner. 400p.
Vettraino, A. M., Natili, G., Anselmi, N. and Vannini, A. (1999). Recent advances in
studies on Phytophthora species associated with Castanea sativa and Quercus
cerris in Italy. In: Phytophthora Diseases of Forest Trees. 1st International JUFRO
Working Party 7.02.09 Meeting. Corvallis, Oregon: Oregon State University.
USA. Hansen, E.M. and Sutton, W. (eds. Pp: 34-36.
Vettraino, A. M., Vannini, A., Anselmi, N. and Fulbright, D.W. (2000). Survey of
Phytophthora species from soil surrounding diseased chesnut species in North
America. In: Phytophthora Diseases of Forest Trees. 1St International IUFRO
Working Party 7.02.09 Meeting. Corvallis, Oregon: Oregon State University.
USA. Hansen, E.M. and Sutton, W. (eds). Pp: 145-147.
Vettraino, A. M., Barzanti, G. P., Bianco, M. C., Ragazzi, A., Capretti, P., Paoletti, E.,
Luisi, N., Anselmi, N. and Vaimini, A. (2002). Ocurrence of Phyt op ht ho ra
species in oak stands in Italy and their association with declining oak trees.
Path. 32: 19-28.
Vettraino, A. M., Belisario, M., Maccaroni, M. and Vannini, A. (2003). Evaluation of
root damage to English walnut caused by five Phytophthora species. Plant
Pathology 52: 491 - 495.
Vettraino, A. M., Morel, 0., Perlerou, C., Robin, C., Diamandis, S. and Vannini, A.
(2005). Ocurrence and distribution of Phytophthora species in European chestnut
stands, and their association with ink disease and crown decline. Eur. J. For. Path.
111: 169-180.
Waterhouse, G. M. (1956). "The genus Phytophthora diagnoses (or descriptions) and
figures from the original papers." CMI. Mycological Papers No. 12. l2Op.
Waterhouse, G. M. (1963). "Key to the species of Phytophthora de Bary." Mycol. Pap.
No. 92: 22p.
62
Waterhouse, G. M. and Waterston, J. M. (1966). C.M.I. Descriptions of Pathogenic Fungi
and Bacteria No. 112, Maps are released periodically by the Commonwealth
Mycological Institute (International Mycological Institute, Bakeham Lane,
Egham, Surrey, United Kingdom).
Werres, S., Marwitz, R., Man In't Veld, W., Dc Cock, A. W. A. M., Bonants, P., De
Weerdt, M., Themann, K., Ilieva, E. and Baayen, R. (2001). Phytophthora
ramorurn sp. nov., a new pathogen on Rhododendron and Viburnum. Mycol. Res.
105(10): 1115-1165.
White, T. J., Bruns, T., Lee, S. and Taylor, J. (1990). Amplification and direct sequencing
of fungal ribosomal RNA genes for phylogenetics Phytophthora cinnamomi. j
PCR Protocols: a Guide to Methods and Applications. M. A. Innis, Geldfand,
D.H., Sninsky, J.J., and White, Ti. (eds). San Diego: Academic Press: 315-321.
Winton, L. M. and Hansen, E. M. (2001). Molecular diagnosis of Phytophthora lateralis
in trees, water, and foliage baits using multiplex polymerase chain reaction.
Path. 31: 275-283.
Zentmyer, G. A., Leary, J. V., Kiure, L. J. and Grantham, G. L. (1976). Variability in
growth of in relation to temperature. Phytopathology 66: 982-986.
63
APPENDICES
64
APPENDIX A
rDNASequence alignments of chinquapin isolates and isolate AG4S clone 2
65
AATACT AATATAC
AATATAC
ATAThC
ATATAC'
I
I
I
8rCTcrTAAc
GG
TCT CrrTTAACT
TCT CTTTTAACT
CTCT CTTTTAACTA
GG
-
90
0917-2
4074
4048
e
TA cAACTTrCA
CTC
CTC
CTC
A
G
TA
TA
'T
TA 7CTC
1
7
i.
H. CAACTTTC*
CAACTTTC
-CAACTTTC
-
7 7 7
7
-I-
e 7
7
Clustal Consensus
'
-'-
*
0917-2
4074
4048
7
AG45___________________
Clustal Consensus
25
26
R7
340
350
0917 -2
4074
4048
AG4 5
Clustal Consensus
360
0917-2
***._____..
Clustal Consensus
410
420
*************** *
440
430
450
460
______
**.* ** **********
470
480
0917-2
ii'jiL1': 1LSjOJ
Clustal Consensus
0917-2
4074
4048
Clustal Consensus
*
*
*
e
e
e
t
7
7
7
jjujjui:
*
*
*
*
*
*
*
APPENDIX B
Pictures of necrotic lesions on ten golden chinquapin trees inoculated with isolates of
Phytophthora cambivora.
gç
Appendix Bi-B4: Bark lesion on chinquapin tree Ml (isolate 4074= Bi;
isolate 09 17-2= B2; isolate 4048= B3 and control= B4) ........................ 67
Appendix B5-B8: Bark lesion on chinquapin tree M2 (isolate 4074= B5;
isolate 09 17-2= B6; isolate 4048= B7 and control= B8) ........................ 68
Appendix B9-B12: Bark lesion on chinquapin tree M3 (isolate 4074 B9;
isolate 0917-2= BlO; isolate 4048= Bil and control B12) ................... 69
Appendix B13-B16: Bark lesion on chinquapin tree M4 (isolate 4074 B13;
isolate 0917-2= B14; isolate 4048= B15 and control= B16) ................... 70
Appendix B17-B20: Bark lesion on chinquapin tree MS (isolate 4074= B17;
isolate 0917-2= B18; isolate 4048= B19 and control B20) ................... 71
Appendix B21-B24: Bark lesion on chinquapin tree Ti (isolate 4074= B21;
isolate 09 17-2= B22; isolate 4048= B23 and control= B24) ................... 72
Appendix B25-B28: Bark lesion on chinquapin tree T2 (isolate 4074= B25;
isolate 09 17-2= B26; isolate 4048= B27 and control= B28) ................... 73
Appendix B29-B32: Bark lesion on chinquapin tree T3 (isolate 4074= B29;
isolate 0917-2= B30; isolate 4048= B31 and control= B32) ................... 74
Appendix B33-B36: Bark lesion on chinquapin tree T4 (isolate 4074= B33;
isolate 0917-2= B34; isolate 4048= B35 and control= B36) ................... 75
Appendix B37-B40: Bark lesion on chinquapin tree T5 (isolate 4074= B37;
isolate 0917-2= B38; isolate 4048= B39 and control= B40) ................... 76
.iç
I&? '-;.
--. :.
-
68
Ttt.
:
r-._
&
-
'h
,2
1jgj
*:;4
p._._
4$
:
:
.'.
!j
j&
r.
B16
'k
70
T
r
if
-4,
-
ii
h
-
- ip I
'p
'Z1
01
72
,;
;
---
i
I
b
7
a-
(41
''.
q,;" 'i b
.
$,..v
.4-
Ø
Il
4'
,'B
75
I
LI
I
U
I
I1:kø Ilp
I'
-
a
77
APPENDIX C
Pictures of golden chinquapin seedlings before and after treatment with isolates of
Phytophthora cambivora under greenhouse conditions
Appendix Cl: Control: chinquapin seedling no.
LA1 .1
before treatment ............. 81
Appendix
C2:
Control: chinquapin seedling no. LA1.1 after treatment ............... 81
Appendix
C3:
Control: chinquapin seedling no.
LB 1.3
before treatment ............. 81
Appendix
C4:
Control: chinquapin seedling no.
LB 1.3
after treatment ............... 81
Appendix C5: Control: chinquapin seedling no. LB2.13 before treatment ............ 81
Appendix C6: Control: chinquapin seedling no. LB2.13 after treatment .............. 81
Appendix C7: Control: chinquapin seedling no. LB2.14 before treatment ............ 81
Appendix C8: Control: chinquapin seedling no. LB2.14 after treatment .............. 81
Appendix C9: Control: chinquapin seedling no. LB2.15 before treatment ............ 81
Appendix ClO: Control: chinquapin seedling no. LB2.15 after treatment ............ 81
Appendix Cli: Control: chinquapin seedling no. LC-2 before treatment ............. 81
Appendix C12: Control: chinquapin seedling no. LC-2 after treatment ............... 81
Appendix C13: Control: chinquapin seedling no. LD1.7 before treatment ............ 81
Appendix C14: Control: chinquapin seedling no. LD1.7 after treatment .............. 81
Appendix C15: Control: chinquapin seedling no. LD2.17 before treatment .......... 81
Appendix C16: Control: chinquapin seedling no. LD2.17 after treatment ............ 81
Appendix C 17: Control: chinquapin seedling no.
LD2. 18
before treatment .......... 82
Appendix
C18:
Control: chinquapin seedling no. LD2.18 after treatment ............ 82
Appendix
C19:
Control: chinquapin seedling no. LD2.19 before treatment .......... 82
Appendix
C20:
Control: chinquapin seedling no. LD2.19 after treatment ............ 82
Appendix
C21:
Inoculated: chinquapin seedling no. LA1.2 before treatment ........... 82
Appendix
C22:
Inoculated: chinquapin seedling no.
LA1 .2
after treatment .......... 82
Appendix
C23:
Inoculated: chinquapin seedling no.
LB1.1
before treatment ......... 82
Appendix
C24:
Inoculated: chinquapin seedling no.
LB 1.1
after treatment ..........
82
Appendix C25: Inoculated: chinquapin seedling no. LB1.2 before treatment ........
82
Appendix C26: Inoculated: chinquapin seedling no. LB1.2 after treatment .......... 82
APPENDIX C (Continued)
Page
Appendix C27: Inoculated: chinquapin seedling no. LB1.4 before treatment ........ 82
Appendix C28: Inoculated: chinquapin seedling no. LB1.4 after treatment .......... 82
Appendix C29: Inoculated: chinquapin seedling no. LB1.5 before treatment ......... 82
Appendix C30: Inoculated: chinquapin seedling no. LB 1.5 after treatment .......... 82
Appendix C3 1: Inoculated: chinquapin seedling no. LB2. 1 before treatment ......... 82
Appendix C32: Inoculated: chinquapin seedling no. LB2.1 after treatment .......... 83
Appendix C33: Inoculated: chinquapin seedling no. LB2.2 before treatment ......... 83
Appendix C34: Inoculated: chinquapin seedling no. LB2.2 after treatment .......... 83
Appendix C35: Inoculated: chinquapin seedling no. LB2.3 before treatment ......... 83
Appendix C36: Inoculated: chinquapin seedling no. LB2.3 after treatment .......... 83
Appendix C37: Inoculated: chinquapin seedling no. LB2.4 before treatment ......... 83
Appendix C38: Inoculated: chinquapin seedling no. LB2.4 after treatment .......... 83
Appendix C39: Inoculated: chinquapin seedling no. LB2.5 before treatment ......... 83
Appendix C40: Inoculated: chinquapin seedling no. LB2.5 after treatment .......... 83
Appendix C41: Inoculated: chinquapin seedling no. LB2.6 before treatment ......... 83
Appendix C42: Inoculated: chinquapin seedling no. LB2.6 after treatment .......... 83
Appendix C43: Inoculated: chinquapin seedling no. LB2.7 before treatment ........... 83
Appendix C44: Inoculated: chinquapin seedling no. LB2.7 after treatment .......... 83
Appendix C45: Inoculated: chinquapin seedling no. LB2.8 before treatment ........ 83
Appendix C46: Inoculated: chinquapin seedling no. LB2.8 after treatment .......... 83
Appendix C47: Inoculated: chinquapin seedling no. LB2.9 before treatment ........ 83
Appendix C48: Inoculated: chinquapin seedling no. LB2.9 after treatment .......... 83
Appendix C49: Inoculated: chinquapin seedling no. LB2.1O before treatment ....... 84
Appendix C50: Inoculated: chinquapin seedling no. LB2.1 1 before treatment ....... 84
Appendix C5 1: Inoculated: chinquapin seedling no. LB2. 11 after treatment ......... 84
Appendix C52: Inoculated: chinquapin seedling no. LB2.12 before treatment ....... 84
Appendix C53: Inoculated: chinquapin seedling no. LB2. 12 after treatment ......... 84
79
APPENDIX C (Continued)
Appendix C54: Inoculated: chinquapin seedling no. LD1(1028) before treatment...84
Appendix C55: Inoculated: chinquapin seedling no. LD1 (1028) after treatment.. ..84
Appendix C56: Inoculated: chinquapin seedling no. LD1.1 before treatment ........ 84
Appendix C57: Inoculated: chinquapin seedling no. LD1.1 after treatment .......... 84
Appendix C58: Inoculated: chinquapin seedling no. LD1.2 before treatment ........ 84
Appendix C59: Inoculated: chinquapin seedling no. LD1.2 after treatment .......... 84
Appendix C60: Inoculated: chinquapin seedling no. LD1.3 before treatment ......... 84
Appendix C61: Inoculated: chinquapin seedling no. LD1 .3 after treatment .......... 84
Appendix C62: Inoculated: chinquapin seedling no. LD1.4 before treatment ......... 84
Appendix C63: Inoculated: chinquapin seedling no. LD1.4 after treatment .......... 84
Appendix C64: Inoculated: chinquapin seedling no. LD1.5 before treatment ........ 85
Appendix C65: Inoculated: chinquapin seedling no. LD1.5 after treatment .......... 85
Appendix C66: Inoculated: chinquapin seedling no. LD1.6 before treatment ........ 85
Appendix C67: Inoculated: chinquapin seedling no. LD1.6 after treatment .......... 85
Appendix C68: Inoculated: chinquapin seedling no. LD2.1 before treatment ......... 85
Appendix C69: Inoculated: chinquapin seedling no. LD2.1 after treatment .......... 85
Appendix C70: Inoculated: chinquapin seedling no. LD2.2 before treatment ......... 85
Appendix C71: Inoculated: chinquapin seedling no. LD2.2 after treatment .......... 85
Appendix C72: Inoculated: chinquapin seedling no. LD2.3 before treatment ........ 85
Appendix C73: Inoculated: chinquapin seedling no. LD2.3 after treatment .......... 85
Appendix C74: Inoculated: chinquapin seedling no. LD2.4 before treatment ........ 85
Appendix C75: Inoculated: chinquapin seedling no. LD2.4 after treatment .......... 85
Appendix C76: Inoculated: chinquapin seedling no. LD2.5 before treatment ........ 85
Appendix C77: Inoculated: chinquapin seedling no. LD2.5 after treatment .......... 85
Appendix C78: Inoculated: chinquapin seedling no. LD2.6 before treatment ........ 85
Appendix C79: Inoculated: chinquapin seedling no. LD2.6 after treatment .......... 85
Appendix C80: Inoculated: chinquapin seedling no. LD2.7 before treatment ........ 86
APPENDIX C (Continued)
Page
Appendix C8 1: Inoculated: chinquapin seedling no. LD2.7 after treatment .......... 86
Appendix C82: Inoculated: chinquapin seedling no. LD2.8 before treatment ........ 86
Appendix C83: Inoculated: chinquapin seedling no. LD2.8 after treatment .......... 86
Appendix C84: Inoculated: chinquapin seedling no. LD2.9 before treatment ........ 86
Appendix C85: Inoculated: chinquapin seedling no. LD2.9 after treatment .......... 86
Appendix C86: Inoculated: chinquapin seedling no. LD2. 10 before treatment ...... 86
Appendix C87: Inoculated: chinquapin seedling no. LD2.10 after treatment ......... 86
Appendix C88: Inoculated: chinquapin seedling no. LD2.1 1 before treatment ...... 86
Appendix C89: Inoculated: chinquapin seedling no. LD2.1 1 after treatment ......... 86
Appendix C90: Inoculated: chinquapin seedling no. LD2.12 before treatment ...... 86
Appendix C91: Inoculated: chinquapin seedling no. LD2.12 after treatment ......... 86
Appendix C92: Inoculated: chinquapin seedling no. LD2.13 before treatment ...... 86
Appendix C93: Inoculated: chinquapin seedling no. LD2.13 after treatment ......... 86
Appendix C94: Inoculated: chinquapin seedling no. LD2.l4before treatment ....... 86
Appendix C95: Inoculated: chinquapin seedling no. LD2.14 after treatment ......... 86
Appendix C96: Inoculated: chinquapin seedling no. LD2.15 before treatment ...... 87
Appendix C97: Inoculated: chinquapin seedling no. LD2. 15 after treatment ......... 87
Appendix C98: Inoculated: chinquapin seedling no. LD2.16 before treatment ...... 87
Appendix C99: Inoculated: chinquapin seedling no. LD2. 16 after treatment ......... 87
1/
.
'I
4:
t
I'
4
-i
81
82
tj
+
DC
85
r;
I
'
87